Avian Blood Flukes (Digenea: Schistosomatidae) in North Dakota Susana Rios

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January 2014 Avian Blood Flukes (Digenea: Schistosomatidae) In North Dakota Susana Rios

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Susana Rios Associate of Science, Johnson College, 2008 Bachelor of Science, Shippensburg University, 2012

A Thesis Submitted to the Graduate Faculty

of the

University of North Dakota

in partial fulfillment of the requirements

for the degree of

Master of Science

Grand Forks, North Dakota December 2014

iii

PERMISSION

Title Avian Blood Flukes (Digenea: Schistosomatidae) in North Dakota

Department Biology

Degree Master of Science

In presenting this thesis in partial fulfillment of the requirements for a graduate degree from the University of North Dakota, I agree that the library of this University shall make it freely available for inspection. I further agree that permission for extensive copying for scholarly purposes may be granted by the professor who supervised my thesis work or, in his absence, by the Chairperson of the department or the dean of the School of Graduate Studies. It is understood that any copying or publication or other use of this thesis or part thereof for financial gain shall not be allowed without my written permission. It is also understood that due recognition shall be given to me and to the University of North Dakota in any scholarly use which may be made of any material in my thesis.

Susana Rios December 1, 2014

TABLE OF CONTENTS

LIST OF FIGURES…………………………………………………..…………………………viii

LIST OF TABLES………………………………………………………………...…..…...……...x

ACKNOWLEDGMENTS………………………………………………………………....….…xii

ABSTRACT………………………………………………………………………………….…xiii

CHAPTER

I. INTRODUCTION………………………………………………………….……..1

Evolution and Phylogeny………………………………………………….2

Morphology…………………………………………………………...... …4

Life Cycle………………………………………………………………….7

Snail Hosts…...………………………………………………..…..….….11

Avian Hosts…...………………………………………………………….11

Avian Health Concerns..……………………………………….……...…12

Effects of Avian Schistosomes in Humans and Other Mammals………………………………………………………..…….….14

Factors Influencing Avian Schistosomatid Diversity and Distribution………...………………………………………………..…...15

Current Status of Avian Schistosomatid Research in the United States……………………………………………………………….…….15

North Dakota as an Ideal Region to Study Avian Schistosomatids……………………………………………………...…..16

Research Objectives………...…………………...…………………...…..17

II. METHODS AND MATERIALS………...……………………………...….……18

Snail Collection……..………………….…………………………..…….18

Snail Husbandry………………………...………………………..…..…..20

Cercarial Screening in Snails…...……………………………….…….…20

Bird Collection…...………………………………………….……..….…22

Bird Dissection Protocol…………...……………………………..….…..22

DNA Extraction…...…………………………………….………...……..23

Polymerase Chain Reaction……………………………………..……….24

DNA Sequencing………...………………………………………………30

Sequence Editing and Alignment…...………………………….…….…..30

Cercarial Morphology…………………………………….…….….…….31

Adult Fluke Morphology……..……………………….………..……..…31

Phylogenetic Analysis……...……………………………….……….…...32

Spatial and Occupancy Analysis of 2013 Snail Data………...... ……33

III. RESULTS……………………………………………………………………..…34

Collected Snails and Avian Schistosomatid Prevalence…….………...…34

Cercariae: Species Identification and Host Associations…..…….….…...40

Adult flukes: Species Identification and Host Associations………....…..42

The Identification of Avian Schistosomatid Species by DNA Sequencing……...…………………………………………..……..…..…45

Intraspecific and Interspecific/Intergeneric Nucleotide Sequence Variation among Collected Avian Schistosomatids………...…...….…..45

Cercarial Morphology…………………………………………..…..……49

Adult Fluke Morphology…..…………………………………….………52

SEM Study of Trichobilharzia spp.………..…………………………………58

Phylogenetic Analysis…...…………………………………………...…..61

Spatial and Occupancy Analysis of 2013 Snail Data……….…………....70

IV. DISCUSSION……………………………………….…………………...………75

Intermediate Snail Hosts……..…………………………………………..75

Definitive Bird Hosts…..…………………………………………….…..78

Sequence Comparison and Phylogenetic Study of Avian Schistosomatids Found in North Dakota…………..…………………….80

Phylogenetic Interrelationships among Avian Schistosomatid Genera…...……………………………………………………………….81

Trichobilharzia……………………………………………….……….….82

Dendritobilharzia……………………………………………….………..83

Austrobilharzia……………………………………………………….….84

Gigantobilharzia……………………………………………...………….84

Cercarial Morphology……………………………………………...…….85

Cercarial Dermatitis in North Dakota……...……………………...……..85

APPENDICES……………………………………………………………………………...……87

A. Map of North Dakota Snail Collection Sites……..………………………..…….88

B. Recorded Data for all Snail Collection Sites…….………………………...…….89

C. Data on Snails Infected with Schistosomatidae Cercariae……..………………...93

REFERENCES………………………………………………………………………………..…97

vii

LIST OF FIGURES

Figure Page

1. Phylogenetic tree of Schistosomatidae based on Bayesian analysis of nuclear 28s rDNA (1200bp)……………...……………………………………………………..…3

2. Male and female avian schistosomatid, Ornithobilharzia sp……………………………...5

3. Variation in size and dimorphism between Schistosomatidae genera (Loker and Brant 2006)……….…………………………………………………..……………….6

4. Life cycle of avian schistosomatids…………………………………………………….....8

5. Phylogeny of intermediate snail hosts and their associated schistosomatidae parasites………………………………………………………………………..…………10

6. Representation of the sampling design used for snail collection. …………………….....19

7. Larval cercarial stage of an avian schistosomatid with identifying morphological features……………………………………………………………………………...……21

8. Ribosomal ITS and 28s nuclear regions showing positions of PCR and sequencing primers………….……………………………………………………….…..26

9. Positions of PCR and sequencing primers in the cox1 mitochondrial gene….………….27

10. All North Dakota snail collection sites from 2013 and the time (month) sites were visited and sampled…………………………………………………………..….…35

11. Map of North Dakota showing 2013 collection sites where AS infected snails were found……………………………………….……..………………………...36

12. Distribution of avian schistosomatid infected Stagnicola snails in North Dakota……………………………………………………...…………………………….37

13. Distribution of avian schistosomatid infected Lymnaea snails in North Dakota……………………………………………………………………………………38

viii

14. Distribution of avian schistosomatid infected Aplexa snails in North Dakota...... ……39

15. Schistosomatidae species found within each snail genus and their prevalence……….…41

16. Morphology of a female Austrobilharzia sp. collected from an Avocet in North Dakota…………………………………………………………………..…………53

17. Morphological comparison between Dendritobilharzia sp. 1 and Dendritobilharzia sp. 2 adult female worms…………………………………………….54

18. Morphological comparison between Dendritobilharzia sp. 1 and Dendritobilharzia sp. 2 adult male worms………………………………………..……..55

19. Complete male Trichobilharzia specimen…………………………………………...…..56

20. Male Trichobilharzia sp. showing the genital opening, testes and terminal end of adult…...………………………………………………………………………….…...57

21. SEM images of a Trichobilharzia szidati cercaria……………………………………….59

22. SEM images of a Trichobilharzia querquedulae adult worm…………………...………60

23. Nuclear 28s rDNA Bayesian tree showing interrelationships among genera of the Schistosomatidae……………………………..…………………………………..…..62

24. A fragment of 28s rDNA Bayesian tree from Fig. 22 showing interrelationships among derived genera of Trichobilharzia, Allobilharzia and Anserobilharzia……….…63

25. Nuclear 28s rDNA Maximum Likelihood tree showing interrelationships among genera of the Schistosomatidae…………………………………………………….…….64

26. A fragment of the 28s rDNA Maximum Likelihood tree from Fig. 24 showing interrelationships among derived genera Trichobilharzia, Allobilharzia and Anserobilharzia……………………………………………………………………..……65

27. Nuclear 28s rDNA Bayesian tree of the derived avian schistosomatid clade…………....66

28. Mitochondrial cox1 Bayesian tree of the genus Trichobilharzia…………………….…..68

29. Mitochondrial cox1 Bayesian tree of Dendritobilharzia specimens collected in North Dakota and one sequence from GenBank……………………………….……..….69

30. Map of North Dakota showing all sites (numbered) where Schistosomatidae infected snails were collected ….……………………………………………..………....88

LIST OF TABLES

Table Page

1. Thermocycling conditions for PCR amplification of 28s rDNA of avian schistosomatid cercariae and adult worms………………………………...……………..28

2. Thermocycling conditions for PCR amplification of mitochondrial cox1 mDNA of avian schistosomatid cercariae and adult worms……………….....…...……..28

3. PCR and sequencing primers for nuclear 28s and mitochondrial cox1 genes used in the study…………………………………………..……………………….……..29

4. Thermocycling conditions for sequencing reactions of nuclear 28s and mitochondrial cox1 DNA for all avian shcistosomatid cercariae and adult worms.….....30

5. Snail genera collected from North Dakota between May-September of 2013 and the number infected with avian schistosomatid cercariae………………….………..40

6. Avian schistosomatids collected from birds in North Dakota……………………….…..43

7. Avian schistosomatid species identified in both the intermediate snail host and the definitive avian host………………………………………………………………….…..44

8. Intraspecific variation of avian schistosomatid species collected in North Dakota based on pairwise sequence comparison………...... ……………………………………..46

9. Interspecific variation of 7 Trichobilharzia species collected in North Dakota based on pairwise sequence comparison……………………………………...……….…47

10. Interspecific variation among members of Austrobilharzia, Dendritobilharzia and Gigantobilharzia in the nuclear 28s (1034 bp) and mitochondrial cox1 (601 bp) genes……………………………………………………………………………48

11. Intergeneric variation of avian schistosomatids collected in North Dakota……………..48

12. Cercarial measurement results for 6 avian schistosomatid species……………………...50

13. Results of ANOVA tests for each cercarial character measured in collected avian schistosomatid cercariae from North Dakota in 2013………………………………..…..51

14. P-value results for cercariae measurements of avian schistosomatids collected in North Dakota……………………………………………………………….…..…..…….51

15. Spatial autocorrelation results for Moran’s I……………………...……………………..72

16. The number of sites for each land cover category in which Lymnaea, Stagnicola and Aplexa snails were present or absent………………………………………………...…..73

17. The number of sites for each ecoregion category in which Lymnaea, Stagnicola and Aplexa snails were present or absent……………………………….…………………....73

18. Fisher’s exact test results showing any significant associations between presence/absence of snail genera (Lymnaea, Stagnicola and Aplexa) and ecoregion or land cover.……………………….…………………………………...…….74

19. Best generalized linear models for the prediction of snail genera presence and infected snail genera presence. …………………………………………….……………74

20. Recorded data for snail collection sites in North Dakota and Minnesota……………....89

21. Data for schistosomatidae infected snails including snail host genus, AS species and sequenced genes…………………………………………...………………..93

ACKNOWLEDGMENTS

I want to thank the members of my advisory committee, Dr. Vasyl Tkach, Dr. Robert

Newman and Dr. Jefferson Vaughan, for their guidance and insight on my research. I would also like to thank the UND faculty and staff for their help and support during my time in the master’s program at the University of North Dakota.

I would like to acknowledge the National Science Foundation, the Joe K. Neel Memorial

Endowment in Limnology and Aquatic Invertebrate Zoology and the UND Department of

Biology APSAC committee for providing funding support to this project. I would also like to acknowledge the Marc Dresden Student Travel Grant from the American Society of

Parasitologists for providing funding to attend and present my research at the 2014 annual meeting in New Orleans.

Special thanks to the people who provided assistance with the collection and care of snails and who also assisted in bird dissections: J. Bell, S. Greiman, E Lawrence K. Neil, R. Nelson,

K.Steffes, K. Patitucci, E. Tkach, M. Tkach and V. Tkach. I would also like to thank the hunters who donated bird specimens during the hunting season: A. Mills, E. Strand D. Anderson, N.

Russert, G. Cain, T. McKenna, M. Flom, S. Greiman, Eric Pulis and P. Burr.

Lastly, I would like to thank my family and friends for their support and encouragement throughout the years.

xii

To my sisters Lissette, Jessie, Evelyn and Jenny.

ABSTRACT

Avian blood flukes are digeneans belonging to the family Schistosomatidae and inhabit the blood circulatory system of birds. Their life cycle includes an intermediate snail host and a definitive bird host. They are found in representatives of several bird orders, but are most prevalent in waterfowl. North Dakota has a large number of wetlands, which provide important breeding sites for many aquatic birds and serve as stopover sites for migratory birds. The abundance and diversity of bird species that congregate in these wetlands ensures optimal conditions for parasite transmission within and among avian species. However, no study on avian blood flukes has ever been conducted in the state and little to no information is available from the surrounding states/provinces as well. The goals of this study were to investigate the diversity, host associations and distribution of avian schistosomatids in North Dakota.

Phylogenetic analysis was also conducted to determine the systematic positioning of blood flukes collected in North Dakota within the Schistosomatidae.

Both the intermediate (snail) and definitive (bird) hosts were surveyed in the state. The larval (cercariae) and adult stages of the parasites were collected from their respective hosts and preserved for morphological and molecular study. Morphological study included measurements of cercariae on temporary mounts, preparing permanent total mounts of adult worms and scanning electron microscopy study of cercariae and adults. Sequences of the partial nuclear ribosomal 28s gene and partial mitochondrial cox1 gene were used in this study for species differentiation and phylogenetic analyses. Sequences obtained in this work were compared with the previously published sequences available in the GenBank database. Intraspecific and

xiii interspecific sequence nucleotide variation was also calculated for all avian schistosomatid species collected in North Dakota. Sequences from North Dakota specimens as well as sequences from all other Schistosomatidae genera available from GenBank were included in a phylogenetic analysis of the Schistosomatidae.

This study focused on the collection of the intermediate snail host to determine the distribution of avian schistosomatid species that may complete their life cycle in the state. Snails were collected from 105 sites throughout the state between May-September of 2013 and screened for the larval stages of parasites. A total of 17,653 snails were collected from six snail genera with an overall infection prevalence of 0.8%. Seven avian schistosomatid species were collected from 4 snail genera. Numerous bird species were also examined and 11 species of avian blood flukes were collected from 22 species of birds. A total of 13 avian schistosomatid species were collected from North Dakota with five species collected from both their intermediate and definitive host. Molecular and morphological analysis also provided evidence for the existence of two Dendritobilharzia species in North America.

These results demonstrated high diversity of avian schistosomatids in North Dakota.

Trichobilharzia was the most speciose genus containing 7 of the 13 avian blood fluke species collected in the state. The majority of infected snails belonged to the genera Stagnicola and

Lymnaea, which were the most heavily sampled snail genera in this study. Further sampling of other snail genera including members of the families Physidae and Planorbidae may reveal the presence of additional avian schistosomatid species circulating in North Dakota. Further sampling of avian hosts, particularly passerines, in North Dakota may also reveal additional species of blood flukes and additional hosts of a widely distributed species Gigantobilharzia huronensis.

xiv

CHAPTER I

INTRODUCTION

Parasites comprise a large portion of all known species (~40%) and it is estimated that there could be 75,000 to 300,000 species parasitizing vertebrates worldwide (Dobson et al 2008).

Birds can harbor many different types of parasites including lice, tapeworms, roundworms and blood flukes (Wobeser 2008). One group of avian parasites that is of particular importance because it can infect both wildlife and people are blood flukes (family Schistosomatidae). Avian blood flukes are a specialized group of parasites that inhabit the circulatory system of their avian hosts. They have been found to infect representatives of multiple bird orders but are most prevalent in waterfowl (Anatidae) (Huffman and Fried 2008). The morbidity/mortality associated with avian schistosomatid infections in wild bird populations is not well understood due to the difficulties in studying wildlife epizootiology, but the pathology caused by avian schistosomatid infection in birds is believed to be comparable with that of the much better studied mammalian schistosomes (Horak et al 2002). Although most parasitic infections do not directly result in the death of their host, they may still reduce host fecundity and affect host survivability, especially in cases where disease and environmental stressors are also present (Wobeser 2008). Recent trends in bird populations have shown a 20-25% reduction in the past 500 years (Sekercioglu et al 2004). Some of the factors attributed to the decline in bird numbers are climate change, habitat loss and fragmentation, and agricultural intensification (Both et al 2006, Newton 2004, Herkert

1994). The combination of environmental stressors with parasitism and disease can ameliorate

1 current declines in bird populations and highlights the importance of studying bird parasites and their associated diversity, transmission, life cycle and distribution (Lafferty and Kuris 1999).

The study of avian schistosomatids is also important for understanding the broader evolutionary history and taxonomic relationships between avian and mammalian schistosomatids within the family Schistosomatidae. Mammalian schistosomes, specifically Schistosoma, have been studied extensively due to the global disease impact associated with human schistosomiasis but information regarding diversity, distribution and host associations for the remaining

Schistosomatidae genera is still lacking from many areas around the world.

This study provides new information on avian schistosomatid diversity and host associations by conducting a survey of avian schistosomatid hosts in North Dakota, one of the most important waterfowl production areas in the United States. North Dakota has a high number of wetlands that serve as breeding grounds and migratory stopover sites for a diverse assemblage of bird species including waterfowl, shorebirds and passerines. This diverse and dense aggregation of birds in shallow, highly productive water bodies provides ideal conditions for avian schistosomatid transmission within and between avian species.

To the best of our knowledge, this is the first study of avian schistosomatids in the state.

The data presented in this thesis provide an overview of the distribution, diversity and host associations of avian schistosomatids in North Dakota as well as their systematic position and phylogenetic affinities.

Evolution and Phylogeny

The family Schistosomatidae includes blood flukes parasitic in birds and mammals. Avian schistosomatids (AS) encompass 10 genera and approximately 67 species (Fig.1). The remaining

Avian genera

Mammalian genera

Figure 1. Phylogenetic tree of Schistosomatidae taxa based on Bayesian analysis of nuclear 28s rDNA (1200bp). Nine identified avian schistosomatid genera (blue boxes) and 4 mammalian genera (red boxes) are included. Three unresolved taxa are also included and are represented by the snail host and location they were collected from. Asterisks denote significant posterior probabilities (>0.95). Figure taken from Brant and Loker 2013

4 genera consist of 30 species of mammalian blood flukes, of which Schistosoma is the most well-known and best studied (Brant and Loker 2013). A phylogenetic study by Snyder (2004), proposed that the Schistosomatidae arose from blood flukes within their sister taxon, the

Spirorchidae. The hypothesis is that a marine turtle blood fluke successfully colonized marine birds and then transitioned to the freshwater life cycle using freshwater avian and snail species

(Snyder 2004). The basal position of the genera Austrobilharzia and Ornithobilharzia (both of which have a marine life cycle) within the Schistosomatidae lends support to this hypothesis as well as the close relationship of marine spirorchid blood flukes to the Schistosomatidae (Snyder

2004).

Morphology

Avian schistosomatids exhibit morphological characteristics typical of most digeneans but also have features unique to their group. They are acoelomates with a dorsoventrally flattened, unsegmented body (Bush et al 2001). Most possess an oral sucker at their anterior end and a ventrally positioned acetabulum that aids in attachment (Fig. 2). They have a syncytial tegument

(outer body covering), which functions in nutrient absorption and protection from environmental conditions during free-living and parasitic life stages. What is unusual about this group is that they are dioecious (separate sexes), whereas most digeneans are hermaphroditic (Khalil 2002).

This has resulted in varying degrees of sexual dimorphism (Fig. 3) for most AS genera (Loker and Brant 2006). Body shape of adult flukes can also vary in shape from filiform to lanceolate as well as in size from a 1.7 mm long in Bilharziella to 57 mm long in Macrobilharzia (Baugh

1963, Fain 1955b).

OS OV

VS SR

VT U VS

SV OS

Figure 2. Male (left) and female (right) avian schistosomatid, Ornithobilharzia sp. OS, oral sucker; SR, seminal receptacle; SV, seminal vesicle; T, testes; U, uterus (with egg); VS, ventral sucker (acetabulum); VT, vitellaria. Figure taken from Faust 1924 (modified).

Figure 3: Variation in size and dimorphism between Schistosomatidae genera (Loker and Brant 2006). Females are on the top and males are on the bottom of each pair. The avian genera are highlighted.

Life Cycle

Avian schistosomatids have a two-host life cycle that includes an intermediate snail host and a definitive avian host (Fig. 4). The typical digenean life cycle usually includes a second intermediate host (three-host life cycle) but this additional step is lost AS because cercariae can directly penetrate the host and cause infection (Rollinson and Simpson 1987). In the three-host life cycle, metacercariae encyst within a secondary intermediate host, (fish, amphibian, crustacean or vegetation) which must then be ingested by the definitive host for infection to occur (Galaktionov and Dobrovolskij 2003). There are free-living and parasitic stages as well as both sexual and asexual reproduction throughout AS development (Horak and Kolarova 2011).

There are also two types of migratory routes AS can utilize within the definitive host (Horak et al. 2002). One of them uses the blood circulatory system (typical of visceral species) whereas the other is through the nervous tissues (typical of nasal forms). It should be noted that only a few species of the genus Trichobilharzia are nasal schistosomatids, while most other schistosomatid genera are visceral forms.

The life cycle of schistosomatids requires an aquatic environment and begins when miracidia hatch from eggs and commence their search for an intermediate snail host. Once they locate a suitable snail, they penetrate its tissue and develop into mother sporocysts. The mother sporocysts then undergo asexual reproduction by producing clusters of germinal cells that develop into daughter sporocysts. The daughter sporocysts migrate to the snail’s digestive organs and produce the next stage, the cercariae. After the cercariae develop, they are shed in cycles dependent on multiple abiotic factors (Galaktionov and Dobrovolskij, 2003). Two of the best known abiotic factors to affect cyclical cercarial shedding are photoperiod and temperature

(Rollinson and Simpson 1987).

Adult

schistosomatid

Sexual reproduction Sexual reproduction

occurs and are and eggs occurs

passed in the feces. passed

Schistosomula travel Schistosomula travel

through blood vessels through

and develop into adults. develop and

Definitive host Definitive

Eggs

C cle

Avian Avian

Cercariae penetrate the penetrate Cercariae

skin of birds.

Micacidia hatch hatch from Micacidia

eggs.

Schistosomatid Life

Cercaria

MIracidia

Snails shed

cercariae. cercariae.

. Life c cle. of avian c schistosomatids. Life

Intermediate host Intermediate

Figure 5 Figure

schistosomatids avian of cycle 4.Life Figure

Once cercariae leave the snail, they are free-swimming and rely solely on their glycogen reserves for energy until they can locate their final avian host. AS cercariae can display different swimming patterns depending on the species, with some found closer to the surface and others in deeper water (Rollinson and Simpson 1987). The swimming behavior and cyclical shedding of cercariae are closely tied to the behavior and activity of their intended definitive host and affects their potential to infect humans. Certain stimuli have been shown to increase swimming activity in cercariae. These stimuli are often indicative of a potential host and include water turbulence, the presence of shadows, and detection of chemicals (ceramides, cholesterol and fatty acids) emitted from the host (Rollinson and Simpson 1987, Haas 2001, Horak and Kolarova 2005,).

When cercariae encounter these stimuli they exhibit continuous swimming with lateral movements introduced to increase their probability of finding a host (Rollinson and Simpson

1987). If a host is not encountered, cercariae resume their standard swimming pattern of upward swimming and passive sinking.

When a definitive host is located, cercariae will attempt to penetrate the epidermis of the potential host. After cercariae enter the epidermis, they undergo a physiological change and transform into schistosomula. The schistosomula migrate through the epidermis until they locate and penetrate a capillary vessel. Once inside the host's vascular system, the schistosomula travel to the lungs and then to the hepatic portal system where they finally develop into adult flukes.

After the adult flukes mature and mate, they travel to the mesenteric veins where they deposit their eggs into the mesenteric venules of the intestine (Platt and Brooks 1997). Dendritobilharzia is an exception to this case and instead utilizes the host’s arterial s stem. The eggs then pass through the walls of the blood vessels, the intestinal wall and into the lumen where they can be excreted with the host’s feces back into the environment. When an influx of water enters the

stosomatidae parasites. Avian schistosomatidae species are sheded are species schistosomatidae Avian parasites. stosomatidae

Figure 5. Ph logen of snail of schi associated Ph 5. logentheir hosts and Figure 2003, al er et modified). (Lock blue. in

10 egg, it induces a mechanical response that breaks open the shell and releases the miracidium into the environment.

Some species of Trichobilharzia have a somewhat different migratory path in the definitive host that involves migration through the bird’s nervous s stem. These species are often referred to as nasal schistosomatids. In this life cycle, schistosomula enter the peripheral nerves after penetrating the skin of birds and migrate through the central nervous system. They move up the spinal cord, into the brain, and then to the nasal blood vessels where they finally mature into adults and release their eggs (Horak et al. 2002). The eggs pass through the nasal blood vessels and into the nasal tissue. Eggs are released back into the environment when the bird places its head in the water (Platt and Brooks 1997).

Snail Hosts

In the United States, both marine and freshwater snails have been implicated in outbreaks of cercarial dermatitis. Freshwater snails are the most common intermediate hosts and include the families Lymnaeidae, Planorbidae, Physidae, Pleuroceridae, Valvatidae and Chilinidae

(Lockyer et al. 2003, Brant and Loker 2013). Overall, snails belonging to 15 families have been reported to be infected with avian schistosomatids (Brant and Loker 2013; Fig. 5). According to

Brant et al. (2006), Lymnaeidae and Physidae are the hosts most frequently used by North

American AS species. In snails, the infection rate is usually between 1-5% (Loy and Haas 2001) but in some locations around the world, it has been estimated to be up to 50% during certain times of the year (Larsen et al 2004, Valdovinos and Balboa 2008).

Avian Hosts

While waterfowl are the most common definitive hosts for AS’s, some species can also parasitize shorebirds, gulls, terns, pelicans, cormorants and songbirds. Several waterfowl species

11 are known to be infected with AS, some of these species include members of the genera Anas

(dabbling ducks including teals, mallards, wigeons, shovelers), Anser (grey geese), Cygnus

(swans), Branta (black geese), Aythya (diving ducks including scaup, redhead, canvasback, ring- necked) Bucephala (bufflehead and goldeneye), Lophodytes (hooded merganser) and Mergus

(mergansers) (Brant and Loker 2013, Vande Vusse 1980).

The prevalence of AS in birds is believed to be much higher compared to the snail intermediate host (Horak and Kolarova 2011). For instance, in one of the studies in France,

60.5% of aquatic birds were infected with AS (Jouet et al 2009) and in Iceland, 35.5% of birds in the orders Gaviiformes, Podicipediformes and Anseriformes were infected (Skirnisson and

Kolarova 2008). Studies in the United States have also found high prevalence within waterfowl including 83.9% prevalence of AS in common mergansers in Flathead Lake, Montana (Loken et al 1995), 24.5% AS prevalence in 46 species of birds (n=378) collected throughout North

America (Brant and Loker 2009b) and 92.8% of three gull species in Connecticut (Barber and

Caira 1995). A study conducted by Brant in 2007 also found that 92% of screened tundra swans from New Mexico and Nevada were infected with AS. Songbirds also may have high prevalence of AS infection; e.g., 60% of red-winged blackbirds in Michigan (Strohm, Blankespoor and

Meier 1981) and 83% of yellow-headed blackbirds in Wisconsin and Michigan (Brackett 1942) were infected with AS.

Avian Health Concerns

The pathological effects of AS on their definitive bird hosts are known to cause inflammation and provoke immune reactions during their migration through tissue. Antigens released from eggs, adults and dead worms can initiate immune responses from the parasitized birds (Horak and Kolarova 2005). The severity of AS infection depends on parasite migratory

12 behavior, parasite load and host immune status (Rau, Bourns and Ellis 1975, Horak et al 2002).

Initial infection occurs when cercariae penetrate the skin and cause minor damage to the epidermal layers. Subsequent phases of infection occur during schistosomula, adult worm and egg migration throughout the visceral organs of the body (Horak and Kolarova 2011). In lung tissue, inflammation, hemorrhage and build-up of scar tissue can result in a compromised respiratory system (Mcmullen and Beaver 1945). Avian schistosomatid adults within the hepatic portal veins can cause liver damage when eggs are deposited into the tissue resulting in a buildup of granulomatous tissue (scar tissue) and inflammation (Mcleod and Little 1942). Infection with blood flukes can also result in blood clots, hemorrhages, and damage to the mesenteric blood vessels (Wojcinski et al 1987) which can increase the susceptibility of infected hosts to other pathogens. Birds with a high parasite load, compromised immune system, or suffering from increased AS pathogenicity as a result of AS migration to other organs are more susceptible to muscle atrophy, reduced kidney and liver function, loss of fat and emaciation (Van Bolhuis et al

2004). The reduction in fitness may affect the overall reproductive potential of their avian host and chronically infected individuals are able to carry and introduce the parasite into new aquatic environments where other individuals can be infected.

In the case of nasal schistosomatids, increased pathogenicity is seen in both specific and non-specific, non-normal avian hosts due to the migration of AS through the CNS. In contrast to visceral schistosomatids in which infection with a low number of AS can be asymptomatic, even a small number of nasal schistosomatids in a host may cause severe neurological symptoms

(Kolarova 2007). Some symptoms of nasal schistosomatid infection can include nasal hemorrhage, balance disorders, paralysis and damage to the spinal cord, brain, ocular nerves and nasal tissue (Horak et al 1999).

Effects of Avian Schistosomes in Humans and Other Mammals

Avian blood flukes can also cause cercarial dermatitis in humans (Brant and Loker 2009a).

This occurs when avian schistosomatid cercariae penetrate human epidermal tissue during their search for an avian host and cause an inflammatory reaction commonly referred to as

‘swimmer’s itch’. The cercariae die in the skin and cause an allergic reaction resulting in a self- limited, non-communicable skin rash. Initially there is a Type I immediate hypersensitivity reaction followed by a late phase inflammatory reaction as a result of the death of cercariae within the skin (Kourilova et al. 2004). No medical treatment is necessary and the rash usually resolves within 1-2 weeks (Kourilova et al. 2004).

Cercarial dermatitis is more commonly reported in the upper Midwest with high numbers seen near the Great Lakes region of the United States (Jarcho and Burkalow 1952.) There have been reports of cercarial dermatitis throughout most of the United States including Pennsylvania,

California and Colorado (Brant et al 2010, Brant and Loker 2009a, Wills et al 1977). Worldwide, cercarial dermatitis has also been reported in Europe, Canada, India, Australia and Japan

(Kolarova 2007).

At this time, very little is known about the survival and migration of AS to other organs of the body in infected mammals. Research has shown migration of some schistosomatid species into the lungs, liver and nervous tissue of mice, rabbits and monkeys (Olivier 1953). It was discovered that three Trichobilharzia species, normally found in waterfowl but known to penetrate the skin of mammals, could migrate to the pulmonary tissue and cause hemorrhage in monkeys (Bacha et al. 1982). One study in Europe found another Trichobilharzia species, T. regenti, in the nervous system of experimentally infected mice. In immunocompromised mice, higher numbers of cercariae were shown to penetrate the skin, migrate to the brain tissue and in

14 some cases, cause paralysis in the infected mouse (Hradkova and Horak 2002). The pathogenicity of AS infection in mammals is still a subject of current research and more studies are needed before we can determine the health risks of AS exposure to mammals.

Factors Influencing Avian Schistosomatid Diversity and Distribution

The distribution of avian schistosomatids is dependent on several abiotic and biotic factors including climate, availability of intermediate and definitive hosts, ecosystem stability, interspecific competition and host immunity. In order for AS to persist in a water body, there must be susceptible intermediate and definitive hosts available to complete the life cycle. An unstable habitat in which rapid changes occur in water level, water quality, vegetation, or species composition can affect AS presence within a water body.

There are also factors that can influence avian schistosomatid diversity including the diversity of intermediate and definitive hosts within a region, their spatial and temporal patterns and the introduction of new potential hosts.

Current Status of Avian Schistosomatid Research in the United States

Currently, the following genera of avian schistosomatids are known to occur in the United

States: Dendritobilharzia, Trichobilharzia, Gigantobilharzia, Allobilharzia, Ornithobilharzia,

Anserobilharzia and Austrobilharzia (Barber and Caira 1995, Lockyer et al 2003, Brant et al

2010). The largest and most broadly distributed genus is Trichobilharzia with 14 reported species. Trichobilharzia physellae is the most common AS species found in snails followed by T. stagnicolae with most cercarial dermatitis outbreaks attributed to one of the two species (Brant and Loker 2009b).

Species misidentification based on morphology alone has become a topic of concern for avian schistosomatids due to insufficiently described specimens from previous research. Before

15 the development of molecular techniques, accurate identification of cercariae and other larval stages was nearly impossible and required experimental infections of intermediate and definitive hosts which was time consuming and costly. This has resulted in incomplete data on the distribution and host associations of avian schistosomatids. Current research utilizes both morphological characteristics and DNA analysis for species identification. This has resulted in the discovery of new species and genera as well as changes to the phylogenetic tree of the

Schistosomatidae (Lockyer et al 2003, Brant and Loker 2009b).

North Dakota as an Ideal Region to Study Avian Schistosomatids

Avian blood flukes have not been previously studied in North Dakota. At the same time, the natural conditions as well as diversity and abundance of aquatic birds in the state makes

North Dakota an ideal region to study avian blood fluke diversity, distribution and host associations. The diversity of bird species that use the migratory stopover sites in North Dakota may also have a large effect on the diversity of schistosomatid species by introducing new species from other regions. Shallow water bodies also facilitate contact between cercariae and their avian hosts and may increase the prevalence of infection.

To the best of our knowledge, there are no records of identified blood flukes or their distribution in North Dakota, although there have been reports of cercarial dermatitis (Jarcho and

Burkalow 1952; Chu 1958). The situation regarding avian schistosomatid knowledge in the neighboring states (South Dakota, Minnesota, Montana) and Manitoba is almost the same, although some sampling was done in Minnesota as part of a molecular phylogenetic study of the genus Trichobilharzia (Brant and Loker 2009b). Considering the abundance of waterfowl and other potential avian hosts of schistosomatids in North Dakota, a relatively diverse fauna and broad distribution of these parasites can be predicted.

Research Objectives

The main objectives of this study are:

 To obtain knowledge of avian schistosomatids, their diversity, distribution and host

associations in North Dakota.

 Create a DNA sequence dataset to enable identification of avian blood flukes found in

North Dakota at any stage of their life cycle.

 Conduct a phylogenetic analysis using newly obtained sequences from North Dakota and

previously published sequences from the GenBank database.

 Perform a spatial analysis of snail distribution in different types of habitats to determine

any distinct patterns and possible correlations with AS infection.

In order to achieve these objectives, snails and birds were collected and examined for the presence of AS cercariae (in snails) or adult blood flukes (in birds) to determine diversity, distribution and host associations in North Dakota. Molecular and morphological techniques were utilized for avian schistosomatid species identification and diversity study. Phylogenetic analysis including North Dakota AS specimens was done in order to determine their relationships with other schistosomatid taxa. Spatial analysis of snail collection sites involved detection of spatial autocorrelation and analysis of snail occupancy.

CHAPTER II

METHODS AND MATERIALS

Snail Collection

The sampling protocol for snail collection was designed so that samples would be a good representation of the different regions in the state, while also maintaining adequate spatial distribution between sites. Locations were selected by organizing the state into a

56 cell grid (approximately 45x30 miles) and selecting at least one area per cell (when possible) which contained several bodies of water in close proximity (Fig. 6). These locations were then designated as areas to visit during a collection trip. Most sites on the eastern half of the state were sampled throughout several day trips between May and

September. Sites in the northern, western and southern part of the state were sampled during four longer (3-5 days) collection trips that occurred in June, July and August of

2013. Water bodies from each of the selected areas were then sampled based on (1) accessibility, (2) water body size, (3) the presence of perennial vegetation (old cattails), and (4) vegetation cover suitable for snail and waterfowl habitation. At each site, GPS coordinates were recorded and pictures of the sampled water bodies were also taken.

Most sites were sampled only once throughout the season. Due to its proximity, the central-eastern part of the state received increased sampling, particularly the region between Devils Lake and Grand Forks. Fewer samples were collected from the southwestern (Badlands) and south-central (Missouri plateau) parts of the state due to the low number of aquatic habitats available in the region. At each site, the sampling protocol

rth rth

n n

Representation the of design sampling used divided Thesnail was state collection. for into with grid a 56 (~45mi cells

hese designated areas were visited and snails were collected if sites met specified criteria. Cell 40 shows a region with Cell criteria. sites region if snails a shows 40 visited areas specified met were and collected were designated hese

a ecoregions (level IV) from the the Hub. (level GIS from ND ecoregions a IV)

Figure 6. Figure bodies selected possible sampling a was as collectio water cell, site. multiple with During at area 1 least xeach 30mi). For t trips, map selected No sampling. of is the for regions multiple in Base were bodies these water locations of densities present, high Dakot

19 required the collection of at least 100 snails of each species per site (if possible) in order for an accurate depiction of AS presence/absence. This sampling size was based on species accumulation curves of cercarial types and AS infection rates of two snail genera obtained from previous snail collections in the central-eastern part of the state.

Snails were collected by gloved hand or dip net. When possible, snails were collected from several locations along the wetland’s shoreline. Water bodies in close proximit (~200ft of each other) were considered one site and snails collected from each water body were pooled and designated as one sample. All collected snails were kept in labeled containers and placed in a cooler to keep snails alive until they could be transported back to the laboratory and examined for AS infection.

Snail Husbandry

Upon collection, snails were housed and screened in the Parasitology Core Facility at the

Department of Biology, University of North Dakota. Snails were rinsed, counted and separated by genus into individual glass jars. Each jar contained 1-4 snails depending on snail size. Jars were filled with specially conditioned tap water to remove chloramine and all jars were cleaned daily or every two days to prevent snail mortality. Snails were fed green leaf lettuce ad lib and maintained at room temperature.

Cercarial Screening in Snails

Two protocols were used to screen snails for cercariae. In the first protocol, snails were kept for 2-5 days and monitored for cercarial shedding once a day for at least two days.

Screening occurred after the snails were exposed to artificial light for ≥ 1 hour. Each jar was screened using a dissecting microscope by examining the bottom, middle and surface water layers for the presence of cercariae. All snails believed to be shedding AS cercariae were

20 dissected using a dissecting microscope to verify AS infection in the snail and to collect larval stages. In the second protocol, snails were not kept in jars or monitored for cercarial shedding.

Instead, all collected snails from a site were immediately dissected and screened for AS infection using the dissecting microscope. The choice of the protocol depended on 1) the duration of the collection trip; longer trips required immediate snail processing because of the difficulty in keeping live snails under field conditions, 2) time constraints; snails were processed as efficiently as possible so that new snails could be collected and screened, and 3) the number of laboratory personnel available for snail processing.

Figure 7. Larval cercarial stage of an avian schistosomatid with identif ing morphological features. Features include a forked tail (A), e espots (B) and knoblike projections on the tips of the tail (C).

Schistosomatid cercariae were identified by using morphological characteristics unique to the group. These morphological characteristics include a forked tail, the presence of eyespots and the presence of knob-like projections on the tips of their tail (Fig. 7). A compound microscope was used in some instances to visualize some of these characteristics. Once identified as AS cercariae, cercariae were preserved in 70-90% ethanol for DNA extraction. Some cercariae were

21 also fixed in hot formalin for scanning electron microscope (SEM) study or photographed using an Olympus BX51 research compound microscope equipped with DIC optics and a digital imaging system utilizing Rincon software.

Bird Collection

Unlike snails, birds were not collected following a specific sampling protocol and were collected opportunistically from 2003-2014. Bird species that are most likely to host AS were targeted when possible. All bird collecting was done according to the obtained federal and state collecting permits as well as approved IACUC protocol. Most birds were collected in the central- eastern part of the state, between Devil’s Lake and Grand Forks, North Dakota. Birds were collected between April and November and included both resident and migratory birds. Most individuals were collected using firearms but some smaller passerines were also captured using mist nets. A number of bird carcasses were provided for examination by local hunters.

Bird Dissection Protocol

All birds were dissected on the day of collection in order to increase the chances of obtaining quality specimens of adult schistosomatids. Birds were identified to species using field guides. During dissection, the liver, kidneys and intestines were removed and kept in a solution of citrated saline to prevent blood clot formation. Sedimentation technique was used to recover blood flukes from kidneys, liver and body wash. The liver and kidneys were each broken up into smaller pieces by gloved hand, and the solution was shaken and allowed to set for 5-10 minutes to allow helminths and tissue time to sink to the bottom of the cylinder. The majority of the supernatant was then discarded and more citrated saline was poured into the cylinder, shaken and allowed to set. This process was repeated until the solution was clear enough to see through under the dissecting microscope. Larger tissue pieces were removed from the solution and small

22 amounts of the solution were poured into a shallow dish and examined under a dissecting microscope. The body cavity of each bird was also rinsed with citrated saline and the resulting body wash also underwent the sedimentation protocol described above. In some cases, it was necessary to pool samples in order to process them in a timely manner. This occurred during the waterfowl hunting season when several waterfowl specimens were donated in a single day.

Pooled samples consisted of birds of the same species that were collected on the same day and from the same site. They were not separated by sex or age.

The mesenteric veins and the blood vessels in the wall of the intestinal tract were carefully examined for the presence of schistosomatid adults. The intestines were removed intact and placed in a large dish containing citrated saline. The mesenteric veins were examined under the dissecting microscope and any worms present were extracted from the blood vessel by excising the blood vessel and pushing the adult flukes out from one end of the vein. Due to the time constraints associated with dissecting multiple birds on the day of collection, it was not always possible to examine the intestines of every bird on instances where several individuals needed to be processed. In such cases, intestines from at least 2-3 birds of each species were examined under the dissecting microscope.

The adult schistosomatids were killed with hot water and preserved in 80% ethanol which permitted both morphological and molecular study. When the number of specimens allowed, some worms/fragments were fixed in 95% ethanol for DNA extraction. Some worms that were intact and in good condition were heat killed and preserved in formalin for SEM study.

DNA Extraction

Schistosomatid cercariae were extracted using two methods. In both cases, 25-30 cercariae were put in an Eppendorf tube. Most of the ethanol was aspirated with a pipette and the

23 specimens were dried for 30 minutes at 60° to remove any remaining ethanol from the tissues of the specimens. Next, 60µl of pure H2O was added to the tube to rehydrate the cercariae. The cercariae were then broken apart by sonication using a UP100H compact ultrasonic processor

(Hielscher USA, Inc., Ringwood, NJ) at 80-100% for 20 seconds. Following sonication, 250µl of

Zymo Cell Lysis Buffer was added to the tubes and the samples were allowed to lyse for at least one hour. In the first method of DNA extraction, the protocol established by Tkach and

Pawlowski (1999) was followed by 1) precipitating DNA with isopropanol for at least two hours or overnight 2) centrifugation and removal of the supernatant 3) rinsing the resulting DNA pellet with 70% ethanol (2x) 4) drying the DNA pellet in a 60° heat block to remove any traces of ethanol. Alternatively, a Zymo micro DNA extraction kit was used according to manufacturer’s instructions. The DNA extraction kit was used for only a few specimens. At the final step of both extraction methods, the DNA was eluted with ≥ 25µl of pure H2O and stored at -20°C.

Adult blood flukes were also extracted using the same protocols as above. DNA was extracted from adults using only partial fragments of adult specimens. In cases of larger specimens like Dendritobilharzia, a posterior or lateral section of the worm was excised and used for DNA analysis. For smaller, filiform specimens like Trichobilharzia, posterior sections of longer worms or fragments of individual worms were used for DNA analysis.

Polymerase Chain Reaction

Following DNA extraction, amplification of a ribosomal and mitochondrial gene was done using polymerase chain reaction (PCR). The two DNA regions chosen for species identification and phylogenetic analysis were nuclear large ribosomal subunit gene (28s) and mitochondrial cytochrome oxidase 1 gene (cox1). In both cases, partial gene sequences were used. It should be mentioned that the sequenced fragment of the cox1 gene included the “barcoding” region widel

24 used for species identification among digeneans and other groups of invertebrates. Fig. 8

(nuclear) and Fig. 9 (mitochondrial) show gene layout and the regions that were amplified and sequenced as well as the positioning of PCR and sequencing primers. The nuclear 28s gene is a component of the large ribosomal subunit and has been used in numerous molecular systematic and phylogenetic studies (Brant and Loker 2009, Snyder 2004). It has both conserved regions as well as variable domains and proved to be useful at different taxonomic levels, from differentiation among congeneric species to phylogenies up to the level of digenean orders.

However, in some digenean lineages the interspecific variability in the 28S gene is not sufficient for reliable species differentiation/identification. Therefore the mitochondrial cox1 gene was chosen as the additional marker characterized by a much higher interspecific variability that allows for differentiation between closely related species.

The PCR protocol included a 20µl reaction consisting of: 10µl of New England Biological

MasterMix Taq Polymerase, 1µl of a forward primer (10pm/µl concentration), 1µl of a reverse primer (10 pm/µl concentration), 1-3 µl of DNA template and enough water to bring the reaction to 20µl. The same protocol was followed for PCR amplification of the mitochondrial cox1 gene.

The PCR conditions for both genes are described in Tables 1 and 2.

In most cases, we have also attempted to amplify the nuclear ribosomal ITS1+5.8S+ITS2 region which has intermediate variability when compared to the 28S and cox1 genes. However, due to difficulty in PCR amplification and DNA sequencing associated with the presence of tandemly arranged repeats and/or multiple copies of the ITS1 region, we were unable to produce an ITS dataset matching that obtained for the 28s and COX1 genes.

1500

1500R1

ITS2

320 BP 320

28s

SCHISTO1300R

D58

5.8S

160BP

900F

ECD2

ITS2

28S

1370BP

2900bp

5.8S

2900bp region that was sequenced. that Positions sequenced. (b) was of PCR and region 2900bp

2600

970 BP 970

ITS1

~750

ITS1

300

SCHISTO100

M18F1

SCHISTO140

ITS5

rimers at the 5’ end of 18S gene and 5.8S gene. (c) Positions of PCR and sequencing primers in the fragment fragment the in primers sequencing PCR Positions and of (c) and gene. 5.8S 18S of gene end 5’ the at rimers

18S

DIGL2

SCHISTO40

18S

ITSF

(a)

(c)

(b)

LSU5

sequencing p sequencing complete ITS and 28s nuclear regions showing 2600 the nuclear 28s and regions complete ITS . this in stud targeted gene 28S of Figure 8. Ribosomal ITS and 28s nuclear regions showing positions of PCR and sequencing primers. showing positions Ribosomal 8. Ribosomal PCR of sequencing regions nuclear 28s (a) and and Figure ITS

COX850R

CO1800RA

1550 1550 BP

COX1 mtDNA COX1

COX555R

COX270F

950BP

COX50F

Figure 9. Positions of PCR and sequencing primers in the cox1 mitochondrial gene. primers Positions 9. gene. of PCR sequencing cox1 the in Figure and mitochondrial

COX1_SCHIST_5 ’

Table 1. Thermoc cling conditions for PCR Table 2. Thermoc cling conditions for PCR amplification of 28s rDNA of avian amplification of cox1 mDNA of avian schistosomatid cercariae and adult worms. schistosomatid cercariae and adult worms. Step Conditions Step Conditions 1 Initial Temp: 94°C 1 Initial Temp: 94°C Denaturation Time: 30 seconds Denaturation Time: 40 seconds 2 Denaturation Temp: 94°C 2 Denaturation Temp: 94°C Time: 30 seconds Time: 30 seconds 3 Annealing Temp: 56°C 3 Annealing Temp: 40°C Time: 40 seconds Time: 45 seconds 4 Extension Temp: 68°C 4 Extension Temp: 68°C Time: 1 minute per kb Time: 1 minute per kb 5 Repeat steps 2-4 X 40 cycles 5 Repeat steps 2-4 X 40 cycles 6 Final Extension Temp: 68°C 6 Final Temp: 68°C Time: 5 minutes Extension Time: 5 minutes

After species identification of all collected specimens, only a few individuals from each species were then chosen for PCR amplification and sequencing of the ITS region.

The PCR products were stored at -20°C and cleaned up using Affymetrix ExoSap enzyme based kit or Zymo DNA Clean and Concentrator column based kit. In cases when multiple bands were present, the Zymo DNA Gel Extraction kit was used to isolate the desired band. All methods were followed according to the manufacturer’s instructions.

Table 3. PCR and sequencing primers for nuclear 28s and mitochondrial cox1 genes used in this study.

PCR amplification and sequencing Primer sequence (5’ – 3’)

NUCLEAR LSU5 TAGGTCGACCCG CTGAAYTTAAGCA 1500R CGAAGTTTCCCTCAGGATAGC 1500R1 GCTACTAGATGGTTCGATTAG ECD2 CCCGTCTTGAAACACGGACCAAG 900F CCGTCTTGAAACACGGACCAAG 300F CAAGTACCGTGAGGGAAAGTTG 300R CAACTTTCCCTCACGGTACTTG SCHISTO100F GTAACTGCGAGTGAACAGGG SCHISTO100R CCCTGTTCAGTCGCAGTTAC SCHISTO1300R GCTCTTGCTCCGCCCCGACG SCHISTO40F GCGGAGGAAAAGAAACTAACAAGG SCHISTO40R CCTTGTTAGTTTCTTTTCCTCCGC DIGL2 AAGCATATCACTAAGCGG DIGL2R CCGCTTAGTGATATGCTT ITSF CGCCCGTCGCTACTACCGATTG ITS5 GGAAGTAAAAGTCGTAACAAGG D58F GCGGTGGATCACTCGGCTCGTG D58R CACGAGCCGAGTGATCCACCGC M18F1 CGTAACAAGGTTTCCGTAG SCHISTO140R CTAAACACCACATTGCCTTGC SCHISTO140F GCAAGGCAATGTGGTGTTTAG MITOCHONDRIAL COX1_SCHISTO5’ TCTTTRGATCATAAGCG COX770R ACCATAAACATATGATG COX555R CCAAATTTWCGATCAAA COX270F GTTTTATATGGARTTGAG COX850R GAAAAAACCTTTATACC CO1800Ra CAACCATAAACATATGATG

DNA Sequencing

The cleaned up PCR products were then cycle-sequenced using Life Technologies BigDye chemistry in a 10µl reaction. The sequencing reaction protocol consisted of: 1-2µl of DNA template, 1.5µl of primer (2pm concentration), 5µl of 5x BigDye sequencing buffer, 1µl of

BigDye Terminator v3.1 Cycle Sequencing Mix and enough water to bring the reaction to 10µl.

Thermocycling conditions are described in Table 4. Sequencing reactions were alcohol- precipitated and sequenced directly on an ABI Prism 3100™ automated capillary sequencer.

Table 4: Thermocycling conditions for sequencing reactions of nuclear 28s and mitochondrial cox1 DNA for all avian schistosomatid cercariae and adult worms. Step Conditions 1 Denaturation Temp: 96°C Time: 15 seconds 2 Annealing Temp: 50°C Time: 10 seconds 3 Extension Temp: 60° Time: 4 minutes 4 Repeat steps 1-3 X 25 cycles

Sequence Editing and Alignment

Sequences were assembled using Sequencher 4.2 software. BioEdit software (Hall 1999) was used for sequence alignment using ClustalW plug-in with default. Subsequent manual adjustment in BioEdit was done when needed. All chromatograms were verified by eye to ensure quality of resulting contigs. Poor quality sequences with background interference were not used.

The nuclear 28s sequences were used for genus identification of collected specimens while the mitochondrial cox1 sequences were used for species identification. Both genes were used for phylogenetic analyses. The ITS regions were not used for phylogenetic analysis due to the

30 presence of tandem repeats. These repeats resulted in difficulty aligning contigs from multiple species and errors in alignment may result in false inferences from any phylogenetic analysis using a misaligned dataset.

Cercarial Morphology

Cercariae preserved in 90% ethanol were rehydrated with water and photographed using a compound microscope utilizing Rincon software. Images were taken of ten cercariae for each haplotype, collected from two different snails (5 from each snail). Measurements were taken from the cercariae images using Rincon software and used for comparison of morphological features between haplotypes. Measurements included body length and body width, oral sucker length and width, ventral sucker length and width, forebody length, tail length, furcae length, and length of the terminal projections of furcae. A few cercariae were also prepared for SEM study.

Fixed specimens in formalin and 70% ethanol were dehydrated in a graded series of ethanol and dried with hexamethyldisilazane (Ted Pella Inc., Redding California) as a transition fluid.

Cercariae were mounted on an aluminum stub using conductive double-sided tape, coated with gold-palladium, and examined with the use of a Hitachi 4700 scanning electron microscope

(Hitachi U.S.A., Mountain View, California) at an accelerating voltage of 5-10kV.

Adult Fluke Morphology

Quality adult worm specimens collected from birds were stained, mounted and photographed. Adult worms preserved in 70% ETOH were first rehydrated in dH2O then stained with alum carmine. Acid ethanol was used as a de-staining fluid to remove any excess stain and worms were then dehydrated using a series of ETOH solutions of increasing concentration.

Afterward, specimens were cleared in clove oil and permanently mounted in xylene-based

Damar gum solution. Some adults were also used for SEM study and were prepared following the same protocol described above for cercariae.

Phylogenetic Analysis

Alignments for analysis were composed of at least two sequences for most of the identified species. When possible at least one sequence was taken from cercariae and one from an adult blood fluke. In addition, sequences from GenBank were also added in the alignment to ensure that as many of the schistosomatid taxa collected from around the world would be represented in the analysis. There were a total of four alignments assembled including 1) nuclear 28s nucleotide sequences representing all species from the derived avian schistosomatid clade (Trichobilharzia,

Gigantobilharzia, Bilharziella, Dendritobilharzia, Allobilharzia, Anserobilharzia ) collected in

North Dakota and available from GenBank, 2) nuclear 28s nucleotide sequences from all species collected in North Dakota and available from GenBank, 3) mitochondrial cox1 nucleotide sequences of Trichobilharzia species from North Dakota and GenBank, and 4) cox1 nucleotide sequences of Dendritobilharzia from North Dakota and GenBank,

All alignments were trimmed to the same length for analyses. jModelTest (Darriba et al

2012, Guindon and Garcuel 2003) and MEGA 6 (Tamura et al 2013) were used to determine the best nucleotide substitution model for each alignment. Phylogenetic analysis using maximum likelihood (ML) were carried out using MEGA 6, and Bayesian inference (BI) using MrBayes

3.2.2 (Huelsenbeck and Ronquist 2001, Ronquist et al 2011). The model GTR+I+G was used for all analyses. Nodal support was estimated by bootstrap (10,000 replicates) and all codon positions were used for ML analyses. For BI (nst=6, rates=invgamma, ngammacat=4) four chains were run simultaneously for 5 x 106 generations and trees were sampled every 1000

32 cycles with the first 1,250 trees discarded as burnin. Trees were visualized and edited using

FigTree v1.4.0 and Adobe Illustrator CS5 software.

Spatial and Occupancy Analysis of 2013 Snail Data

The data gathered from snail collections in North Dakota (2013) was also used to assess spatial patterns of snail and parasite distribution. I tested for the presence of distinct snail distribution patterns through spatial autocorrelation and occupancy in relation to ecological context using logistic regression (Guisan et al 2002). For spatial autocorrelation, SAM (Spatial

Analysis in Macroecology) v4.0 (Rangel et al 2010) was used to calculate Moran’s I for sites where Lymnaea, Stagnicola or Aplexa snails were collected and for sites where AS infected

Lymnaea, AS infected Stagnicola or AS infected Aplexa snails were collected. Distribution data on Physa, Helisoma and Promenetus snails were not included in this analysis due to small sample size and lack of AS infections. Physa were infected in only two sites while no AS infections have been detected in the latter two genera. For spatial autocorrelation analysis, the

105 sites were separated into 12 distance classes containing equal numbers of pairs and 500 permutations were run.

Snail occurrence based on ecoregion and land cover was performed using Fisher’s exact- test. Land cover and ecoregion for each collection site was determined by plotting sites on

Omernik ecoregion (2013) and LandCover (2013) maps using ArcGIS 10.2.1. Logistic regression, using R 3.1.1, was applied to determine the best model for predicting snail presence.

The possible predictor variables included land cover, ecoregion, collection date, latitude and longitude. The same analysis was performed for schistosomatid occurrence in which the presence of an infected snail genus was the response variable.

CHAPTER III

RESULTS

Collected Snails and Avian Schistosomatid Prevalence

A total of 17, 653 snails were collected between May – September of 2013 from 105 sites throughout North Dakota (Fig. 10, Appendix A and B). Prolonged cold temperatures during the spring of 2013 delayed the onset of snail sampling for the season and only one site was sampled in late May. The months with the highest number of sampled sites were July (40 sites) and

August (29 sites). Eighteen sites were sampled in June and 17 sites were sampled in September.

Thirty six out of 105 sites were positive for avian schistosomatid infected snails (Fig. 11). A large portion of the sampled sites (91%) were located in the Northwestern Glaciated Plains and

Northern Glaciated Plains ecoregions (Fig. 11).

Collected snails represent 6 genera and 3 families (Table 5). There were no avian schistosomatid infected snails from the genus Helisoma and Promenetus. The avian schistosomatid prevalence rate within snail genera was highest in Lymnaea (1.7%) and Aplexa snails (2.9%), while both Stagnicola and Aplexa had AS prevalence rate < 0.5%. A total of 134

(0.8%) snails were shedding cercariae. The distribution of infected Stagnicola, Lymnaea and

Aplexa snails are shown in Figs. 12–14.

In addition, 195 snails were collected from 5 sites in eastern-central Minnesota with 2 sites positive for avian schistosomatid infected snails. They included one Physa snail and two

Lymnaea snails. Snails shedding mammalian schistosomatids (Schistosomatium douthitti) were

May

July

June August

September

Month site was was site Month sampled

All North Dakota snail collection sites from 2013 and the time period (b month) sites were visited and sampled. sampled. visited month) and were sites (b period time the and 2013 sites from Dakota collection North All snail

Figure 10. Figure the map from ecoregions Omernik North of is the (level Dakota IV) Base ear. the during onl visited once Most were sites Hub. NDGIS

not collected not AS infected AS snails

AS infected AS snails collected

Lake Lake

Plain

Agassiz Agassiz

Northwestern

Glaciated Plains

Northern

Glaciated Plains

nails were collected. Base map of Omernik ecoregions (level II) taken from ND GIS hub. hub. from taken GIS ND Omernik ecoregions of map (level Base II) collected. were nails

Map of North Dakota showing 2013 collection sites where AS infected snails were represent snails found.were circles infected AS Red Map collection 2013 showing sites where North of Dakota

Northwestern Great Plains

Figure 11. Figure no avian sites where represent circles Black collected. avianwere snails schistosomatid sites infected where collection s schistosomatidinfected

snails snails

Stagnicola collected and and collected AS with infected

Stagnicola not but collected AS with infected

Collection site Collection

an schistosomatid cercaria (blue (blue circles). schistosomatid cercaria an

snails collected (black circles), 2) collection site collection 2) circles), (black collected snails

snails in North Dakota. Snail collection sites are in snails collection Snail Dakota. sites are North

Stagnicola

snails were collected and at least one was infected was one avi with least and collected at snailswere

snails were collected but none were infected with avian schistosomatids ( ½ black, ½ blue circles), 3) infected circles), blue ½ avian with none but snails schistosomatidswere black, ½ ( collected were

Stagnicola

Figure 11. Distributionof schistosomatid avian infected 11. Figure dividedinto categories three 1) that site sampled was but no where sitewhere

snails snails

Lymnaea

Lymnaea Lymnaea and collected AS with infected

Lymnaea Lymnaea not but collected AS with infected

No collected snails

snails in North Dakota. Snail collection sites are divided into into divided are sites collection Snail Dakota. in snails North

stosomatids (½ black, ½ red circles n=36), where site red ½ circles stosomatids3) black, (½

snails collected (black circles), 2) collection site where site where 2) circles), (black collected snails collection

Lymnaea

Figure 12. Distribution of avian schistosomatid infected infected schistosomatid of avian Distribution 12. Figure three categories 1) no but sampled was categories that site three but collected with infected schi were avian none were snails n=25). circles (red infected schistosomatid cercariae was one avian with least and collected at were snails

snails

snails snails

Aplexa

collected and and collected Aplexa Aplexa AS with infected

infected with AS with infected Aplexa Aplexa not but collected

Aplexa

No collected

snails in North Dakota. Snail collection sites are divided into divided North in snails collection Snail sites Dakota. are

Aplexa

snails collected (black circles), 2) collection site where 2) circles), collection where site (black collected snails

schistosomatid cercariae (green circles, n=7). circles, (green cercariae schistosomatid

Aplexa

were collected and at least one was infected with avian with infected was least one at and collected were Figure 13. Distribution of avian schistosomatid infected Distribution 13. of infected schistosomatid Figure avian 1) no but sampled was categories that site three n=14), circles, infected where site 3) green ½ avian with none but schistosomatidswere black, (½ collected were

39 also collected in North Dakota and included four Stagnicola snails and 9 Lymnaea snails. These snails are not included in Table 5. Differentiation of mammalian and avian cercariae was based on DNA sequencing.

Table 5. Snail genera collected from North Dakota between Ma -September of 2013 and the number infected with avian schistosomatid cercariae. Snail Family Snail Genus Total Collected N. infected with AS cercariae Lymnaeidae Stagnicola 10893 38 (0.4%) Lymnaea 4260 73 (1.7%) Physidae Physa 1157 2 (0.2%) Aplexa 732 21 (2.9%) Planorbidae Helisoma 495 0 Promenetus 116 0 Total 17653 134 (0.8%)

Cercariae: Species Identification and Host Associations

A total of 8 schistosomatid species were identified from cercariae collected from snails, seven AS species and one mammalian schistosomatid (Fig. 15, Appendix C). Stagnicola served as intermediate hosts for the highest number of schistosomatid species with a total of 6 species detected form this genus. Lymnaea snails had four species detected with only one positive snail for Trichobilharzia sp. A and Trichobilharzia sp. E. AS species were identified to species by matching the newly obtained sequences to sequences already available in the GenBank database.

Five out of the 7 sequenced AS species, matched GenBank sequences of identified species. Two

AS species did not have close matches to any already identified species using GenBank sequences and are designated as Trichobilharzia sp. 2 and Trichobilharzia sp. 4. Trichobilharzia szidati was the most prevalent avian schistosomatid species collected from 71 Lymnaea snails.

The least prevalent was Trichobilharzia sp. 2 with only two infected snails collected (both from the same site). The mammalian schistosomatid, Schistosomatium douthitti, was found to utilize both Lymnaea and Stagnicola snails as intermediate hosts and the avian schistosomatid species,

Gigantobilharzia huronensis, was also found in two snail genera, Physa and Aplexa. The AS species Trichobilharzia szidati, Trichobilharzia sp. A and Trichobilharzia sp. E were found within two snail genera as well, however, the low prevalence in one of the two snail genera indicate that snail misidentification may be more likely.

Adult Flukes: Species Identification and Host Associations

Avian schistosomatids were found in 72 birds collected in North Dakota between 2003 and

2014. In the fall of 2013, there were nine instances in which samples had to be pooled. For these samples, they are described as coming from a single bird since it is unknown how many total birds within the pooled sample may have been infected. Mallards, Blue-winged teals and

Northern shovelers were the bird species with the highest numbers of individuals examined.

There were 11 AS species recovered from 22 species of birds collected in North Dakota (Table

6, Appendix D).

Trichobilharzia was the most diverse AS genus in birds with 5 species obtained. One

Trichobilharzia species, Trichobilharzia sp. 4, did not have a close match to any identified species with sequences in the GenBank database. All sequenced Dendritobilharzia specimens had identical nuclear 28s rDNA sequences that matched Dendritobilharzia pulverulenta sequences available from GenBank. However, they formed two distinct groups when the mitochondrial cox1 gene sequences were analyzed. These two Dendritobilharzia groups were named Dendritobilharzia sp. 1 and Dendritobilharzia sp. 2. Within the genus Gigantobilharzia,

Table 6. Avian schistosomatids collected from birds in North Dakota. Number of Schistosomatidae species Bird species infected birds Allobilharzia visceralis Cygnus columbianus (Tundra swan) 1 Branta canadensis (Canada goose) 1 Austrobilharzia sp. Recurvirostra americana (Avocet) 1 Limosa fedoa (Marbled godwit) 1 Dendritobilharzia sp. 1 Anas clypeata (Northern shoveler) 1 Anas platyrhynchos (Mallard) 3 Anas discors (Blue-winged teal) 2 Aythya valisineria (Canvasback) 1 Dendritobilharzia sp. 2 Anas platyrhynchos (Mallard) 1 Anas clypeata (Northern shoveler) 1 Anas strepera (Gadwall) 3 Bucephala albeola (Bufflehead) 1 Aythya americana (Redhead) 1 Dendritobilharzia sp. Aythya affinis (Lesser scaup) 1 Gavial immer (Common loon) 1 Anas clypeata (Northern shoveler) 1 Anas platyrhynchos (Mallard) 1 Aix sponsa (Wood duck) 1 Gigantobilharzia huronensis Quiscalus quiscula (Grackle) 5 Xanthocephalus xanthocephalus (Yellow-headed 1 blackbird) Agelaius phoeniceus (Red-winged blackbird) 3 Molothrus ater (Brown-headed cowbird) 1 Gigantobilharzia sp. Gallinago gallinago (Common snipe) 1 Trichobilharzia sp. E Anas crecca (Green-winged teal) 2 Trichobilharzia querquedulae Anas discors (Blue-winged teal) 6 Anas strepera (Gadwall) 1 Anas clypeata (Northern shoveler) 4 Trichobilharzia sp.4 Anas clypeata (Northern shoveler) 2 Anas discors (Blue-winged teal) 1 Trichobilharzia sp. A Anas strepera (Gadwall) 2 Trichobilharzia physellae Anas platyrhynchos (Mallard) 2 Aythya collaris (Ring-necked duck) 2 Aythya affinis (Lesser scaup) 2 Trichobilharzia sp. Aythya collaris (Ring-necked duck) 2 Aythya americana (Redhead) 2 Anas discors (Blue-winged teal) 1 Anas clypeata (Northern shoveler) 6 Bucephala clangula (Common goldeneye) 1 Aix sponsa (Wood duck) 1 Anas platyrhynchos (Mallard) 1

43 all sequences matched the Gigantobilharzia huronensis from GenBank except for one adult fluke recovered from a Common snipe. The closest match was to the genus Gigantobilharzia, so this specimen was named Gigantobilharzia sp. A specimen collected from a Marbled godwit also did not match any available GenBank sequence. The closest match was to the genus Austrobilharzia, therefore this specimen is referred to as Austrobilharzia sp. The second Austrobilharzia specimen was a single complete worm that was stained and mounted instead of undergoing DNA extraction. Since no DNA sequences are available for this specimen, it is also labeled as

Austrobilharzia sp.

There are 5 avian schistosomatid species for which both the cercarial and larval stages were recovered from the intermediate snail and definitive bird host (Table 7).

Table 7. Avian schistosomatid species identified in both the intermediate snail host and the definitive avian host.

Avian schistosomatid species Intermediate snail host Definitive avian host Trichobilharzia sp. E Stagnicola Anas crecca (Green-winged teal) Trichobilharzia querquedulae Aplexa Anas discors (Blue-winged teal) Anas strepera (Gadwall) Anas clypeata (Northern shoveler) Trichobilharzia sp. 4 Stagnicola Anas clypeata (Northern shoveler) Anas discors (Blue-winged teal) Trichobilharzia sp. A Stagnicola Anas strepera (Gadwall) Gigantobilharzia huronensis Aplexa Quiscalus quiscula (Grackle) Physa Xanthocephalus xanthocephalus (Yellow-headed blackbird) Agelaius phoeniceus (Red- winged blackbird) Molothrus ater (Brown-headed cowbird)

The Identification of Avian Schistosomatid Species by DNA Sequencing

All collected samples of cercariae from snails and the majority of samples of adult blood flukes from birds underwent DNA extraction and PCR amplification. In some instances, quality

DNA sequences could only be obtained for either the mitochondrial cox1 or the nuclear 28s large ribosomal subunit. Due to the relatively conserved nature of the nuclear 28s gene, differentiation of some species (e.g., Trichobilharzia querquedulae, Trichobilharzia sp. A and Trichobilharzia sp. 4) was only possible using the cox1 gene. As a result, 8 cercarial and 14 adult blood fluke samples of Trichobilharzia could only be identified to the genus level. The same scenario occurred among Dendritobilharzia specimens in which differentiation was only possible using the mitochondrial cox1 gene. Dendritobilharzia specimens that did not produce quality cox1 sequences (n=5) were only identified to genus.

Intraspecific and Interspecific/Intergeneric Nucleotide Sequence Variation

among Collected Avian Schistosomatids

The intraspecific nucleotide variation for all collected AS species is shown in Table 8 for both the nuclear 28s (1034bp) and mitochondrial cox1 (578 bp) genes. No intraspecific sequence variation was detected in the nuclear 28s gene for all collected species except Gigantobilharzia huronensis and Trichobilharzia querquedulae. The latter two species both had one sequence containing one base pair difference. The intraspecific nucleotide variations, within the mitochondrial cox1 gene, were all higher but averages were still below 1%.

The interspecific nucleotide variation for AS species collected in North Dakota is shown in

Table 9 (Trichobilharzia species only) and Table 10 (all other collected AS species). When comparing interspecific nucleotide variation for the 28s gene, there was an average variation of

1.21% for all collected species. The average interspecific variation for the mitochondrial cox1

Table 8. Intraspecific variation of avian schistosomatid species collected in North Dakota based on pairwise sequence comparison. The nuclear 28s (1034) and mitochondrial cox1 (578bp) genes were used for comparisons. Avian schistosomatid # Sequences Intraspecific # Sequences Intraspecific species for 28s variation for for cox1 variation for 28S, % cox1, % min-max (avg) min-max (avg) Allobilharzia visceralis 2 0 1 - Austrobilharzia sp. 1 - 1 - Dendritobilharzia sp.1 7 0 5 0-0.99 (0.47) Dendritobilharzia sp. 2 4 0 5 0.20-1.38 (0.83) Gigantobilharzia 5 0-0.09 6 0-0.60 huronensis (0.03) (0.20) Gigantobilharzia sp. 2 1 - 1 - Trichobilharzia sp. E 2 0 5 0.17-2.00 (0.77) Trichobilharzia sp. 2 2 0 2 0.33 Trichobilharzia 3 0-0.08 5 0-1.50 querquedulae (0.06) (0.67) Trichobilharzia sp. 4 4 0 5 0.33-0.83 (0.63) Trichobilharzia sp. A 2 0 6 0-1.33 (0.72) Trichobilharzia szidati 5 0 10 0-1.50 (0.50) Trichobilharzia 2 0 6 0-0.60 physellae (0.27)

gene was 11.64%. To demonstrate the significant level of divergence between the

Austrobilharzia sp. collected in North Dakota and Austrobilharzia variglandis, a sequence of A. variglandis from GenBank was used for comparison (Table 10).

Table 11 shows intergeneric variation in studied genera. The 28s intergeneric nucleotide variation was lowest between Dendritobilharzia and Gigantobilharzia and highest between

Allobilharzia and Austrobilharzia. The cox1 intergeneric nucleotide variation was lowest between the genera Allobilharzia and Trichobilharzia and highest between the genera

Dendritobilharzia and Gigantobilharzia. The cox1 sequence obtained from the Austrobilharzia sp. collected in North Dakota was 200 bp shorter than sequences from all other genera so it was

0.29

), 2.71 2.03 0.19 0.19 0.87

****

(0.23)

physellae

. .

0.19

sp. 4 (4 4 sp.

variation in the in variation

0.97

ise sequence sequence ise

szidati

1.84 1.35 0.87 0.87

the the

****

9.82

11.15

(0.90)

. .

(10.51)

0.87

sp. A sp.

12.65 13.31

. .

sequences used for each species species each for used sequences Trichobilharzia

- -

0.01

2.71 2.03

based on pairw on based

****

(0.05)

(12.06) (13.13)

11.31 12.98

(3 Tricho

/cox1

sp. 4 sp.

11.65 9.82

11.31

- -

0.1

2.71 2.03

****

(0.03) (9.68)

(10.41) (11.05)

9.48

querquedulae

9.48

10.48

. . Tricho

Variability is shown as % of the sequence length. The min The length. sequence the of as % shown is Variability

). ).

2.80 2.13 5.99 11.98 9.82

- - - - -

10.48 (

sp. 2 (2 2 sp.

****

9.53)

(2.74) (2.06) (5.36) (9.48)

(11.38)

species collected in North Dakota inNorth collected species

2.71 2.03 4.99 9.32

querquedulae

10.82

8.82

. .

physellae

. .

sp. sp. 2

/10

12.98 13.31 14.14 13.48 14.31

. .

- - - - -

Trichobilharzia

2.22

****

Trichobilharzia

(12.70) (12.93) (13.53) (13.28) (14.23)

12.48 12.48 12.81 12.81 14.14

Tricho

szidati

. .

sp. E (2 E sp. ),

sp. E sp.

12.15 12.82 13.98 14.98 11.81 14.64

------

variation of 7 7 of variation

****

(12.01) (12.51) (13.50) (14.34) (11.38) (14.50)

fic

11.98 12.15 12.65 13.48 10.48 14.31

Trich

avg) are followed by average in parentheses. Only one number is shown where no variability was was variability no where shown is number one Only in parentheses. average by are followed avg)

sp. A (2 A sp.

. The variation in the nuclear 28s partial gene (1034bp) is shown above the diagonal and and diagonal the above shown is (1034bp) gene partial 28s the nuclear in variation The .

Trichobilharzia

. .

. Interspeci .

d: d:

szidati

sp. 2 sp. 4 sp.

sp. E sp. sp.

. .

physellae

species

T .

querquedulae

Trichobilharzia Trichobilharzia Trichobilharzia Trichobilharzia Trichobilharzia

comparison 28s of number The diagonal. the below isshown (601bp) gene cox1 mitochondrial include Trichobilharzia ( and values max detected. Table

Table 10. Interspecific variation among members of Austrobilharzia, Dendritobilharzia and Gigantobilharzia in the nuclear 28s (1034 bp) and mitochondrial cox1 (601 bp) genes. The number of sequences used for 28s/cox1 comparison included: Austrobilharzia sp. (1/1), Dendritobilharzia sp. 1 (7/5), Dendritobilharzia sp. 2 (4/5), Gigantobilharzia huronensis (5/6) and Gigantobilharzia sp. 2 (1/1). Values are shown as percentages. Avian schistosomatid species compared Interspecific Interspecific variation for 28S, variation for cox1, min-max min-max (avg) (avg) Austrobilharzia sp. -Austrobilharzia variglandis * 0.30 13.50

Dendritobilharzia sp. 1 -Dendritobilharzia sp. 2 0 6.71-7.30 (7.02) Gigantobilharzia huronensis -Gigantobilharzia sp. 2 0-0.37 10.88-11.33 (0.14) (11.28) *An Austrobilharzia variglandis sequence from GenBank was used for comparison.

Table 11. Intergeneric variation of avian schistosomatids collected in North Dakota. The intergeneric variation in the nuclear 28s partial gene (1098bp) is shown above the diagonal and intergeneric variation in the mitochondrial cox1 gene (578bp) is shown below the diagonal. Variability is shown as % of the sequence length. The min-max values are followed by average in parentheses.

Genera Allobilharzia Austrobilharzia Dendritobilharzia Gigantobilharzia Trichobilharzia Allobilharzia 11.34 5.28 5.28-5.65 4.10-4.92 ***** (5.52) (4.61) Austrobilharzia - 9.53 9.62-9.98 9.35-10.16 ***** (9.86) (9.82) Dendritobilharzia 19.72-20.42 - 2.55-2.73 2.91-4.46 ***** (20.18) (2.67) (3.56) Gigantobilharzia 17.88-20.31 - 20.59-22.67 2.72-4.54 ***** (18.18) (22.07) (3.68) Trichobilharzia 13.74-17.04 - 16.96-21.45 15.80-20.83 ***** (15.42) (18.69) (18.34)

excluded from intergeneric comparisons to preserve sequence length of other genera (Table 11).

Intergeneric variation for comparisons with Austrobilharzia sp. were calculated using only 325 bp and the averages are as follows: Allobilharzia (21.54%), Dendritobilharzia (23.32%),

Gigantobilharzia (25.35%) and Trichobilharzia (21.18%).

Cercarial Morphology

The measurements for AS cercaria and their variance are shown in Table 12. Not all measurements were taken for all collected species because only a few infected snails were collected for some AS species. There were also lower numbers of cercariae collected from smaller sized snails like Aplexa, and Physa. Trichobilharzia querquedulae was the only AS species for which no measurements were taken due to low numbers of cercaria available after

DNA extraction (Aplexa is the intermediate snail host) and Trichobilharzia sp. 2 only had one snail, with 5 cercariae measured. Difficulty visualizing the oral and ventral suckers for all measured cercariae also produced low, unreliable measurements in some instances, therefore the measurements for oral sucker length/width, and ventral sucker width are not reported.

One way ANOVA tests were performed for all cercarial features measured and values are reported in Table 13. Forebody length was the only feature measured that did not show any significant difference between avian schistosomatid species, F (3.25) = 0.82, p = 0.54. Post hoc

T-tests were then conducted for all other measured cercarial features and results are shown in

Table 14. To correct for the problem of performing multiple T-tests simultaneously, the

Bonferroni correction method was utilized to adjust the P-value significance cutoff from 0.05 to

0.0006. Most comparisons between Gigantobilharzia huronensis and Trichobilharzia species resulted in significant differences between cercarial measurements. There were few significant differences in morphological measurements when Trichobilharzia species were compared to each other. Comparisons of tail length and body width between Trichobilharzia sp. 2 and other

Trichobilharzia species had the highest number of significant p-values (7). There were also a few

(3) significant p-values when Trichobilharzia sp. 4 was compared to other Trichobilharzia species.

6.1 4.4 6.6 5.0 7.6 4.3 8.7 8.8

15.0 16.2 13.6 11.6 15.2 22.5 17.6 11.4 11.0 12.2

2.5 9.4 2.4 2.6 1.9 1.7 2.0

30.7 20.6 17.3 19.6 17.0 42.7 11.9 16.0 16.7 21.2 34.5

StDev

20.7 243.3 365.3 416.2 322.0 358.0 373.4 227.8 225.1 216.8 271.7 265.4 19.5 18.4 17.5 18.5

Max

15.6

122.9

- - - - -

------

- -

7.5

Min

13.3 10.5 14.5 12.6 12.4

77.9

151.0 301.3 370.0 249.9 308.9 253.7 174.3 202.5 170.5 208.4 144.9

16.8 11.0 15.0 16.2 15.4 16.6

192.8 337.3 390.0 294.5 344.3 316.8 101.1 209.7 220.7 195.9 235.9 230.0

Median

Avg

16.5 10.8 14.7 16.4 15.6 16.3

189.3 337.9 393.0 294.8 338.3 314.6 102.0 210.1 216.6 191.3 240.9 227.1

sp. A sp. sp. E sp. 2 sp. 4 sp. A sp. E sp. 2 sp. 4 sp. A sp. E sp. 2 sp. 4 sp.

Tail length Tail

Furcae length Furcae

Trichobilharzia Gigantobilharzia huronensis Gigantobilharzia Trichobilharzia Trichobilharzia Trichobilharzia Trichobilharzia szidati Trichobilharzia huronensis Gigantobilharzia Trichobilharzia Trichobilharzia Trichobilharzia Trichobilharzia szidati Trichobilharzia projections furcae of Length huronensis Gigantobilharzia Trichobilharzia Trichobilharzia Trichobilharzia szidati Trichobilharzia

5.5 7.8 5.8 5.9 9.0 7.6 2.5 3.1 6.6 7.1 6.1 5.4 5.6 3.9

10.8 12.7 10.9 33.9 12.2 17.1 13.0 12.6

- -

sp. 2. 2. sp.

1.8 8.7 2.8 4.5 7.1 4.6 2.4 1.0 5.3 3.9 3.0

23.9 11.8 22.7 19.0 16.4 27.2 24.7 12.8 64.9 11.2 12.7

StDev

255.1 226.5 326.7 352.5 305.6 360.0 350.7 85.1 92.6 77.5 75.6 222.3 237.1 216.4 233.8 22.8 26.6 36.7 36.0 27.7

Max

------

76.94

56.068.6

Min

71. 52.4 85.6 62.1 55.7 18.4 25.2 26.2 26.2 21.1

199.6 195.8 263.0 305.0 253.1 269.6 284.2 192.8 145.3 194.7 211.6

Trichobilharzia

6 avian schistosomatid species. Measurements were taken for ten cercariae of each species (5 cercariae cercariae (5 species each of cercariae ten for taken were Measurements species. schistosomatid avian 6

- -

73.8 67.7 91.4 67.6 62.4 65.3 18.9 25.9 30.3 29.0 21.8

210.2 212.4 289.1 316.0 271.1 306.4 323.3 209.8 191.2 210.3 233.3

Median

- -

Avg

74.0 68.3 90.4 67.9 64.9 63.6 20.0 25.9 31.1 29.7 23.5

222.1 212.8 291.8 324.5 278.3 302.1 322.7 208.7 191.2 207.1 226.2

sp. A sp. sp. E sp. 2 sp. 4 sp. A sp. E sp. 2 sp. 4 sp. A sp. E sp. 2 sp. 4 sp. E sp. 2 sp. 4 sp. A sp.

arzia szidati arzia

Cercarial measurement results results measurement Cercarial

. . width Body

Bodylength

Forebody length Forebody

Ventral sucker length sucker Ventral

x 2 snails). Only 5 cercariae were measured from from measured were cercariae 5 Only snails). 2 x Trichobilharzia Gigantobilharzia huronensis Gigantobilharzia Trichobilharzia Trichobilharzia Trichobilharzia Trichobilharzia szidati Trichobilharzia huronensis Gigantobilharzia Trichobilharzia Trichobilharzia Trichobilharzia Trichobilharzia szidati Trichobilharzia huronensis Gigantobilharzia Trichobilharzia Trichobilharzia Trichobilharzia szidati Trichobilharzia huronensis Gigantobilharzia Trichobilharzia Trichobilharzia Trichobilharzia Trichobilharzia Trichobilh Table 12 Table

Table 13. Results of ANOVA tests for each cercarial character measured in collected avian schistosomatid cercariae from North Dakota in 2013. P-values below 0.05 are in bold.

df between df within Cercariae feature F F crit P-value species species

Tail length 5 48 59.6 2.41 2.0x10-19

Furcae length 5 48 58.2 2.41 3.2x10-19

Furcae projection length 5 46 8.47 2.42 9.6x10-6

Body length 5 48 34.58 2.41 8.2x10-15

Body width 5 48 18.43 2.41 3.6x10-10

Ventral sucker length 4 13 5.59 3.18 0.007 Forebody length 4 12 0.82 3.26 .54

Table 14. P-value results for cercariae measurements of avian schistosomatids collected in North Dakota. T-tests were used to compare 6 cercariae features measured for 6 avian schistosomatid species. P-values below 0.0006 (6.0x10-4) are considered significant and are in bold. Species being compared Body Body Ventral Tail Furcae Furcae sucker length length projection width length length length

Gigantobilharzia huronensis x Trichobilharzia sp. E 0.068 1.7x10-7 - 1.1x10-9 5.4x10-12 0.003 Gigantobilharzia huronensis x Trichobilharzia sp. 2 3.0x10-5 3.2x10-5 - 7.0x10-10 1.6x10-9 6.2x10-4 Gigantobilharzia huronensis x Trichobilharzia sp. 4 0.002 1.5x10-8 - 1.4x10-7 2.2x10-10 2.1x10-4 Gigantobilharzia huronensis x Trichobilharzia sp. A 0.003 5.1x10-7 - 2.2x10-9 3.8x10-11 9.3x10-5 Gigantobilharzia huronensis x Trichobilharzia szidati 7.1x10-5 8.4x10-8 - 6.8x10-7 1.5x10-6 8.0x10-5 Trichobilharzia sp. E x Trichobilharzia sp. 2 8.7x10-6 0.015 0.039 3.4x10-4 0.338 0.189 Trichobilharzia sp. E x Trichobilharzia sp. 4 0.922 0.146 0.052 1.5x10-5 0.020 0.380 Trichobilharzia sp. E x Trichobilharzia sp. A 0.357 0.369 0.005 0.970 0.002 0.141 Trichobilharzia sp. E x Trichobilharzia szidati 0.163 0.012 0.135 0.279 0.202 0.155 Trichobilharzia sp. 2 x Trichobilharzia sp. 4 5.5x10-8 0.002 0.231 3.6x10-6 0.003 0.488 Trichobilharzia sp. 2 x Trichobilharzia sp. A 4.5x10-7 0.182 0.200 8.2x10-4 0.018 1.869 Trichobilharzia sp. 2 x Trichobilharzia szidati 1.6x10-8 0.884 0.168 5.4x10-4 0.411 0.946 Trichobilharzia sp. 4 x Trichobilharzia sp. A 0.269 0.032 0.718 5.2x10-5 2.1x10-5 0.401 Trichobilharzia sp. 4 x Trichobilharzia szidati 0.054 4.6x10-4 0.116 0.093 0.016 0.459 Trichobilahrzia sp. A x Trichobilharzia szidati 0.643 0.102 0.023 0.258 0.322 0.869

Adult Fluke Morphology

There were four species for which complete adult worms were collected, stained and mounted for morphological study. The Austrobilharzia sp. collected from an Avocet is shown in

Fig 16. Only a single worm was collected so no molecular data is available for this

Austrobilharzia sp.

Multiple Dendritobilharzia sp. 1 and Dendritobilharzia sp. 2 specimens were stained and mounted and consisted of both genetically marked and unmarked adult worms. In order to identify morphologic features that could be used to differentiate between the two proposed

Dendritobilharzia species, genetically marked individuals were examined and are shown in Fig.

17 (female) and 18 (male). Two morphological features that were found to vary between the two species included the position of the genital pore in relation to the cecal bifurcation in male

Dendritobilharzia specimens. In genetically marked Dendritobilharzia sp. 1 specimens, the ceca loop rejoined anterior to the testes and the location of the genital pore was adjacent to the ceca bifurcation point (Fig. 18a). In genetically marked Dendritobilharzia sp. 2 specimens, the ceca loop continued into the testes where it rejoined posterior to the first pair of testes (Fig. 18b). The location of the genital pore occurred at the midpoint region of the cecal loop. Another morphological characteristic that appeared to be a possible identifying feature of each species was their overall body shape with Dendritobilharzia sp. 1 specimens having a wider body shape and Dendritobilharzia sp. 2 specimens having a more slender body shape (Fig 17a,b and Fig

18a,b).

There were few complete adult worms of Trichobilharzia due to the difficulty in extracting complete worms from the mesenteric blood vessels of birds and low intensity of infection in most birds. Trichobilharzia querquedulae was the only Trichobilharzia species in which

O S OV

OV OV

C C B

a b c Figure 16. Morphology of a female Austrobilharzia sp. collected from an Avocet in North Dakota. (a) Complete specimen, (b) ovary and (c) ceca. OS, oral sucker; OV, ovary; C, ceca.

53 a EO b EO c UT

UT UT CE OV OV

SR SR SR VT CB VT d

e OV

Figure 17. Morphological comparison between Dendritobilharzia sp. 1 (a) and Dendritobilharzia sp. 2 (b) adult female worms. Reproductive structures are shown in (c). (d) and (e) show an empty and full seminal receptacle respectively. CB, cecal bifurcation; CE, ceca; EO, eosophatus; OV, ovary; SR seminal receptacle; UT, uterus; VT, vitellaria.

54 a b c EO EO SV

SV GP GP SV

CB GP CB

TE TE CB d

Figure 18. Morphological comparison between Dendritobilharzia sp. 1 (a) and Dendritoblharzia sp. 2 (b) adult male worms. A closer view of the positioning of the genital pore and cecal bifurcation are shown for Dendritobilharzia sp. 1 (c) and Dendritobilharzia sp. 2 (d). CB, cecal bifurcation; EO, eosophagus; GP, genital pore; SV, seminal vesicle; TE, testes.

a OS b c OE OS VS SV

SV d VS

e SV GC

Figure 19. Complete male Trichobilharzia specimen (a). Anterior portion of a female Trichobilharzia specimen (b)showing the ventral sucker. Anterior portion of a male Trichobilharzia specimen (c) Seminal vesicle of a male Trichobilharzia specimen (d and e) and the beginning of the g naecophoric canal (e). GC,g naecophoric canal; OE, oesophagus; OS, oral sucker; SV, seminal vesicle; VS, ventral sucker.

a GP

b TE

c TE

Figure 20. Male Trichobilharzia specimen showing the genital opening (a) testes (b) and terminal end of adult (c). GP, genital pore; TE, testes.

57 complete adult worms were collected in sufficient numbers for morphological study. A few of these worms were used for SEM study. One complete adult worm collected from a shoveler was stained and mounted (Fig. 19 and 20). No molecular data is available for this specimen and it was only identified to genus.

SEM Study of Trichobilharzia spp.

We have studied some larval and adult Trichobilharzia specimens under scanning electron microscope. SEM study of Trichobilharzia szidati cercariae allowed for a detailed observation of the external morphology. Fig. 21a shows the body of the cercaria with a lateral view of the ventral sucker. The fin folds of the bifurcated tail and the projections on the tips of the furcae are shown in Fig. 21b, c. Minute spines were found distributed not only throughout the body of the cercaria (Fig. 21d), but also on the surface of the tail and even furcae (Fig. 21c). Sensory papillae were seen on the body of the cercariae primarily in the zone of suckers (Fig. 21e, f).

SEM study of an adult Trichobilharzia querquedulae (Fig. 22) allowed us to examine the external structure of both the oral and ventral suckers, the tegumental spination as well as the external structure of the gynecophoric canal in the male worm (Fig 22). Sensory papillae were also seen distributed throughout the tegumental surface of the adult worm (Fig. 22c).

VS FF

A B FP C

SP S

D E F

Figure 21. SEM images of a Trichoiblharzia szidati cercaria. A) Body of cercaria with ventral (VS) and oral sucker (OS). B) Tail furcae (FC) and furcae projections (FP) on the tail tips. C) Closer view of spines (S) and fin fold (FF) present on tail. D) Spines (S) on body of cercariae. E) Ventral and F) lateral view of sensory papillae (SP) on body of cercaria.

pillae (PP) on on (PP) pillae

B C

adult worm. A) Partial anterior portion of an adult male with visible oral (OS) and (OS) oral with visible male adult ofan portion anterior Partial A) adultworm.

querquedulae

Trichoiblharzia

ventral (VS) sucker and the anterior portion of the gynecophoric canal (GC). B) Closer view of the ventral sucker (VS). C) Pa C) (VS). sucker ventral the of view Closer B) (GC). canal thegynecophoric of portion anterior the and sucker (VS) ventral surface. tegumental Figure 22. SEM images of a a of images SEM 22. Figure

Phylogenetic Analysis

A total of 8 trees were generated using maximum likelihood and Bayesian analysis for the

4 alignments described in Chapter II. The nuclear 28s rDNA alignment of Schistosomatidae genera included 91 sequences (1100 bp) with 26 sequences obtained from avian schistosomatid specimens collected in North Dakota (13 species, 5 genera). Sequences also included 1) one sequence of a mammalian schistosomatid species collected from a lymnaeid snail in North

Dakota, Schistosomatium douthitti, 2) one sequence of a Gigantobilharzia species collected from a Sandwich tern (Thalasseus sandvicensis) in Mississippi, 3) one sequence from a Bilharziella polonica obtained from a Pochard (Aythia ferina) collected in the Ukraine and 4) two

Macrobilharzia macrobilharzia sequences obtained from Anhingas (Anhinga anhinga) collected in Mississippi. The remaining sequences were taken from the GenBank database. A total of 14

Schistosomatidae genera and >30 recognized species are represented as well as 16 avian schistosomatid sequences with unknown systematic placement. Several regions throughout the world are represented including North America, South America, Asia, Africa, Europe and

Australia with the majority of sequences obtained from snails and birds collected in North

America.

Bayesian and Maximum Likelihood both produced trees of similar topologies with

Bayesian analysis resulting in more strongly supported nodes. The Bayesian and Maximum

Likelihood nuclear 28s rDNA trees are each separated into two figures due to better visualization of relationships between taxa (Figs. 23-26). The systematic positions of Trichobilharzia,

Allobilharzia and Anserobilharzia are shown in Fig. 24 (Bayesian analysis) and Fig. 26

(Maximum Likelihood analysis). There is a difference in topology between the two trees regarding the placement of Trichobilharzia physellae within the Trichobilharzia sp. A,

Fig 23. Nuclear 28s rDNA Bayesian tree showing interrelationships among genera of the Schistosomatidae. Sequences obtained in this stud are marked with “ND”. Posterior probability values are shown above the internodes. Collapsed part of the tree is presented on Figure 23. Genera are color coded to show monophyletic and non-monophyletic lineages. Posterior probability values are shown. - 62

Figure 24. A fragment of the 28S Bayesisn tree from Fig. 22 showing interrelationships among derived genera Trichobilharzia, Allobilharzia and Anserobilharzia. Trichobilharzia species are color coded to show monophyletic and non-monophyletic lineages. Posterior probability values are shown at the internodes.

Figure 25. Nuclear 28s rDNA Maximum likelihood tree showing interrelationships among genera of the Schistosomatidae. Sequences obtained in this stud are marked with “ND”. Bootstrap values are shown above the internodes. Collapsed part of the tree is presented on Figure 25. Genera are color coded to show monophyletic and non-monophyletic lineages. Bootstrap values are shown above the internodes.

Figure 26. A fragment of the 28s rDNA Maximum Likelihood tree from Fig. 24 showing interrelationships among derived genera Trichobilharzia, Allobilharzia and Anserobilharzia. Trichobilharzia species are color coded to show monophyletic and non- monophyletic lineages. Bootstrap values are shown at the internodes.

Figure 27. Nuclear 28s rDNA Bayesian tree of the derived avian schistosomatid clade. Genera are color coded to show monophyletic and non-monophyletic lineages. Posterior probabilities are shown.

Trichobilharzia querquedulae, Trichobilharzia sp. 4 and Trichobilharzia franki clade. There is also a difference in tree topology regarding the positioning of the genera Allobilharzia and

Anserobilharzia in relation to Trichobilharzia. These two genera are positioned within the

Trichobilharzia clade in the Bayesian tree but are positioned as a sister taxon to Trichobilharzia in the Maximum Likelihood tree. The remaining Schistosomatidae genera are shown in Fig 23

(Bayesian analysis), and Fig. 25 (Maximum likelihood). The tree topology is almost identical in both trees except for slight changes in relative position of weakly supported branches that are treated as polytomies.

The phylogenetic tree on Fig. 26 is based on Bayesian analysis of 28s sequences of members of the derived avian schistosomatid clade consisting of Trichobilharzia, Allobilharzia,

Gigantobilharzia, Dendritobilharzia, Bilharziella and Anserobilharzia with Schistosomatium used as an outgroup. The taxon set in this tree is compatible with that used by Brant et al. (2013) and Brant and Loker (2013). The topology in this tree is almost identical to the 28s Bayesian tree containing all Schistosomatidae genera (Fig. 23) except for slight changes in the polytomies containing Gigantobilharzia, Dendritobilharzia and avian schistosomatids from Argentina.

Increased variability within the mitochondrial cox1 gene allowed for finer resolution within genera, however, this high nucleotide variability also produced phylogenetic trees with low node support for both Maximum likelihood and Bayesian analyses when multiple genera were included in the alignments. The mitochondrial cox1 phylogenetic trees of the derived avian schistosomatid clade (Trichobilharzia, Gigantobilharzia, Bilharziella, Dendritobilharzia,

Allobilharzia and Anserobilharzia) produced trees with multiple polytomies and low branch support (trees not shown). Only the Bayesian analyses of alignments containing one genera (570 bp) produced tree topologies with moderate support and they are depicted in Figs. 28 and 29.

Figure 28. Mitochondrial cox1 Bayesian tree of the genus

Figure 28. Mitochondrial cox1 Bayesian tree of the genus Trichobilharzia. Species are color coded to demonstrate monophyletic and paraphyletic grouping within species. Species with only one representative are in black. Posterior probabilities are shown.

sp. 2 sequences are highlighted in red. in highlighted are sequences 2 sp.

specimens collected in North Dakota and one sequence from GenBank. GenBank. from sequence one and Dakota North in collected specimens

Dendritobilharzia

sp. 1 sequences are highlighted in blue and inblue highlighted are sequences 1 sp.

re 29. Mitochondrial cox1 Bayesian tree of of tree Bayesian cox1 Mitochondrial re 29. Figu Dendritobilharzia

Fig. 28 shows the systematic positioning between Trichobilharzia species. There is separation between the species Trichobilharzia querquedulae, Trichobilharzia sp. 4,

Trichobilharzia franki, Trichobilharzia physellae and Trichobilharzia sp. A into separate taxa.

These species were grouped into the same clade during Bayesian and Maximum Likelihood analyses of the nuclear 28s rDNA gene. Trichobilharzia szidati/Trichobilharzia ocellata formed a strongly supported (99%) clade. However, the interrelationships among taxa belonging to this clade were not well resolved. Sequences from specimens collected in Europe and Russia (with the exception of a single US specimen) and published as T. ocellata and T. szidati formed a polytomy. In contrary, all specimens of T. szidati collected by us in North Dakota formed a

100% supported clade. This suggests a possibility that the present T. szidati may in fact contain more than one species. Allobilharzia and Anserobilharzia are also positioned within the

Trichobilharzia clade in this Maximum Likelihood tree of the mitochondrial cox1 gene.

The final tree (Fig. 29) was generated using Bayesian methods and includes only

Dendritobilharzia mitochondrial cox1 gene sequences from specimens collected in the United

States. The proposed species, Dendritobilharzia sp. 1 and Dendritobilharzia sp. 2, separate into two distinct clades supporting the results of the pairwise sequence comparisons presented above

(Table 10).

Spatial and Occupancy Analysis of 2013 Snail Data

Spatial autocorrelation was only seen in sites where Lymnaea snails were present, indicating non-independence for these sites (Table 15). Sites within 25 km of each other showed the highest level of spatial autocorrelation (Moran’s I=0.348, p-value=0.002). The distance class which included sites within 65 km of each other also showed a significant level of spatial autocorrelation (Moran’s I= 0.146, p-value=0.006). No spatial autocorrelation was detected in

70 sites where Stagnicola and Aplexa snails were collected or sites where AS infected Lymnaea, AS infected Stagnicola or AS infected Aplexa snails were collected.

Sites were divided into 7 land cover (Table 16) and 4 ecoregions (Table 17). Because of small sample sizes, three land cover (deciduous forest, n=3; developed, n=1; fallow crop, n=2) and one ecoregion category (Lake Agassiz Plain, n=2) were removed from the analysis. The

Bonferroni correction method was used to adjust the significance cutoff (exact-p) from 0.05 to

0.0013 (ecoregion) and 0.017 (land cover). The Fisher’s exact-test showed a significant association between land cover and the presence of Lymnaea snails, exact p=0.0005 (2-tailed test) (Table 18). Herbaceous wetlands had the highest occupancy rate for Lymnaea snails (Table

16). There was no significant difference in the occupancy rate of Stagnicola and Aplexa snails based on land cover or ecoregion.

Logistic regression (Table 19) revealed land cover (herbaceous wetlands, grass pastures and open water) and latitude/longitude to be the best predictors for the presence of Lymnaea snails at a site. Date of sampling was also shown to be a good predictor for the presence of

Aplexa snails. No models were found to be good predictors for Stagnicola presence and for the presence of AS infected Lymnaea snails, AS infected Stagnicola snails and AS infected Aplexa snails.

Table 15. Spatial autocorrelation results for Moran’s I. This analysis was performed on snail collection sites where Lymnaea, Stagnicola, or Aplexa snails were collected as well as sites where AS infected Lymnaea, AS infected Stagnicola and AS infected Aplexa snails were collected. The 105 sites were divided into 12 distance classes with equal number of pairs (count=908). Significance was tested using 500 permutations. P-values < 0.05 are in bold. LymPres StagPres AplPres DistCntr (km) Moran's I P-value Moran's I P-value Moran's I P-value 24.381 0.348 0.002 0.029 0.482 0.038 0.368 64.739 0.146 0.006 0.017 0.726 -0.014 0.732 94.745 0.064 0.134 -0.049 0.278 -0.031 0.494 120.624 -0.058 0.188 -0.044 0.308 0.016 0.722 143.569 -0.12 0.012 -0.032 0.454 -0.023 0.608 164.827 -0.103 0.024 -0.004 0.936 -0.009 0.858 188.173 -0.018 0.668 -0.034 0.426 -0.056 0.206 216.157 -0.108 0.028 -0.057 0.24 0.013 0.758 243.721 -0.121 0.006 0.04 0.365 -0.058 0.156 275.975 -0.204 0.002 0.018 0.668 -0.017 0.682 329.247 0.19 0.002 -0.008 0.826 0.052 0.252 494.422 -0.126 0.006 0.007 0.826 -0.014 0.638 Lym_AS_Pres Stag_As_Pres Apl_AS_Pres DistCntr (km) Moran's I P-value Moran's I P-value Moran's I P-value 24.381 -0.026 0.564 0.028 0.49 0.029 0.528 64.739 -0.012 0.764 -0.025 0.556 -0.022 0.588 94.745 -0.035 0.412 -0.005 0.908 -0.048 0.262 120.624 0.004 0.938 0.018 0.662 -0.089 0.048 143.569 0.016 0.674 -0.076 0.088 0.038 0.358 164.827 -0.071 0.1 -0.077 0.084 0.048 0.29 188.173 0.037 0.408 0.002 0.968 -0.01 0.836 216.157 -0.042 0.362 0.06 0.162 0.033 0.46 243.721 -0.048 0.254 0.035 0.428 -0.053 0.162 275.975 0.079 0.062 -0.112 0.02 -0.077 0.062 329.247 -0.016 0.706 0.038 0.36 0.033 0.416 494.422 0.002 0.926 0.002 0.936 0.004 0.87

Table 16. The number of sites for each land cover category in which Lymnaea, Stagnicola and Aplexa snails were present or absent. n=the number of sites for each category. Land Crop Deciduous Developed Fallow Grass Herbaceous Open Cover (n=24) forest (n=1) Crop Pasture wetland Water (n=3) (n=2) (n=36) (n=29) (n=10) Lymnaea 12 2 0 1 15 26 5 Present Lymnaea 12 1 1 1 21 3 5 Absent Stagnicola 20 3 1 2 33 25 8 Present Stagnicola 4 0 0 0 3 4 2 Absent Aplexa 4 0 0 0 10 6 1 Present Aplexa 20 3 1 2 26 23 9 Absent

Table 17. The number of sites for each ecoregion category in which Lymnaea, Stagnicola and Aplexa snails were present and absent. n= the number of sites for each ecoregion category. Ecoregion Northwestern Northwestern Northern Lake Agassiz Glaciated Plains Great Plains Glaciated Plain (n=34) (n=7) Plains (n=2) (n=62) Lymnaea 16 4 40 1 Present Lymnaea 18 3 22 1 Absent Stagnicola 30 7 53 2 Present Stagnicola 4 0 9 0 Absent Aplexa 7 0 14 0 Present Aplexa 27 7 48 2 Absent

Table 18. Fisher’s exact test results showing any significant associations between presence/absence of snail genera (Lymnaea, Stagnicola and Aplexa) and ecoregion or land cover. Exact p-values < 0.013 (ecoregion) and <0.017 (landcover) are considered significant (2-tailed) and are in bold. LandCover Ecoregion Df Exact P df Exact P LymPres 3 0.0005 2 0.298 StagPres 3 0.761 2 0.641 AplPres 3 0.569 2 0.463

Table 19. Best generalized linear models for the prediction of snail genera presence and infected snail genera presence. LCgp=grass pasture, LChw, herbaceous wetland, LCow, open water, Long=longitude, Jdate=date Lymnaea AS Stagnicola AS Aplexa AS presence infected presence infected presence infected Lymnaea Stagnicola Aplexa presence presence presence Best LCgp, p=0.003 - - - Jdate, - model LChw, p<.001 p<.001 LCow, p=0.13 Long, p<.001 Null 130.725 - - - 93.068 - deviance Residual 90.315 - - - 77.337 - deviance AIC 102.32 - - - 81.837 -

CHAPTER IV

DISCUSSION

This study was primarily focused on the examination of intermediate snail hosts because it would better depict the regional diversity and distribution of avian schistosomatids in North

Dakota. Since these aquatic snails are confined to a water body, the assumption can be made that transmission of the AS species found in snails occurs locally. Avian schistosomatids found in birds on the other hand, cannot be assumed to complete their life cycle within North Dakota because the examined birds included adult migratory birds. This has resulted in finding AS species for which adult flukes do not have a corresponding cercarial match from snails. The opposite case, in which AS cercariae are collected but their corresponding adult fluke stage is not found within birds also occurred. In the latter case, two different scenarios are possible. We either did not collect an appropriate bird host species or we could have collected the hosts, but did not find a parasite due to small sample size.

Intermediate Snail Hosts

The AS prevalence in snails collected in North Dakota is similar to what was found in previous studies which have shown an average AS prevalence between 1-5% (Loy and Haas

2001). The majority of our sampling effort was focused on the collection of Stagnicola and

Lymnaea snails. Fewer numbers were collected for other snail genera; therefore, comparisons of

AS prevalence in snail genera may be biased.

Stagnicola and Lymnaea were the most common snail genera in North Dakota wetlands.

Their large size and high numbers make them an ideal intermediate host in these aquatic habitats.

Other snail genera/species that were not sampled or sampled insufficiently (e. g., Gyraulus) may also play an important role in AS transmission. Since the majority of wetlands that were sampled in 2013 were of similar size and vegetative structure, it is likely that other snail taxa are present in other types of aquatic habitats. Planorbidae and Physidae snails, which were not collected in large numbers during this study, are known to serve as intermediate hosts for multiple avian schistosomatid species, including Dendritobilharzia, Trichobilharzia, and Anserobilharzia species (Brant 2009a, 2013). These snail genera are present in North Dakota (Fuliss et. al. 1999) and are likely to serve as intermediate hosts for other schistosomatid species not found in this study.

Spatial analysis of sampled sites from 2013 detected the presence of spatial autocorrelation in sites where Lymnaea snails were collected, indicating non-independence for these sites. This could indicate a clumped distribution pattern for water bodies containing Lymnaea snails or a clumped distribution of the habitat conditions that are favored by Lymnaea snails. However, these results could also be an artifact of snail sampling methods because sampling was more extensive in areas with high water body density. Occupancy analysis of collected snail genera also revealed a correlation between Lymnaea snail presence and herbaceous wetlands and geographic location (latitude, longitude). Increased sampling in the northeastern part of North

Dakota and the selection of similar types of wetlands most likely influenced these results.

Lymnaea snails were typically found in wetlands with moderate to high vegetative cover while

Stagnicola snails were found in the majority of water bodies sampled (89% of sampled sites).

The low sample size of Aplexa snails collected, relative to the area that was sampled, was not sufficient to produce any reliable results on Aplexa distribution in North Dakota.

Only one avian schistosomatid species, namely Gigantobilharzia huronensis, was found to infect two different mollusk genera, Stagnicola and Lymnaea (Fig 14). In contrast, all the remaining AS species (including all Trichobilharzia) collected in our study were specific to their intermediate host species. Stagnicola had the highest diversity of AS among all snail genera in the region followed by Aplexa while Lymnaea hosted only one AS species (Fig. 14).

No avian schistosomatid infected snails were found in the southwestern part of the state. This region, known as the Badlands, has a semi-arid climate with low densities of water bodies. The inability to collect AS infected snails in this region is most likely due to the limited sampling conducted in the area. Physid snails were the most commonly collected snail genera in this region. Future sampling may reveal AS species utilizing these snails as intermediate hosts in this part of the state.

In our study, the prevalence of AS infections in snails was highest in mid- to late summer.

Multiple factors play a role in determining the time period for which AS prevalence is highest.

Long winters can delay and shorten the amount of time wetlands are available for snails and birds, and long periods of drought or periods with increased precipitation will also affect AS prevalence in intermediate snail hosts. Relative importance of these factors may vary from year to year; however, multi-year observations would be necessary to adequately study this question.

Another factor that could also affect AS prevalence in snails is competition with other parasites for the invertebrate hosts (Bush et al 2001). One potential competitor is Schistosomatium douthitti, a mammalian schistosomatid. This species was collected from both Stagnicola and

Lymnaea snails, two snail genera that were also utilized by avian schistosomatids. In some localities that were repeatedly sampled, eastern –central North Dakota, they account for almost half of the Schistosomatidae infected Lymnaeid snails collected at these sites. Other cercarial

77 types that were also seen to utilize these snail genera included xiphidiocercariae, echinostome and monostome cercariae. For instance, redial stages of echinostomatids are well known to directly feed on larval stages of other digeneans including schistosomatids (Lie 1973).

In this stud , cercariae were taken directl from the snail’s hepatopancreas. This method would have included immature cercariae and would therefore not produce reliable results for avian schistosomatid cercarial measurements. Even though an attempt was made to only include fully formed cercariae, a preferable method would have been the use of free-swimming cercariae from the water column. This would have ensured that cercariae used for measurements were fully formed. Previous studies however, have concluded that using cercarial morphological characteristics is not a reliable method for species differentiation (Brant and Loker 2009a). Size of cercariae can vary depending on snail size and snail species infected. There may also be environmental factors that affect cercarial size causing variation between geographic locations.

Definitive Bird Hosts

A total of 8 avian schistosomatid species were recovered from waterfowl species, 2 from shorebirds and 1 avian schistosomatid species from passerines. Despite a reasonably good coverage of aquatic bird diversity in the study, increased sampling may reveal additional avian schistosomatid species from birds in North Dakota. One of the most intriguing results was the fact that Trichobilharzia szidati was the AS species most prevalent in snails, however, none were recovered from birds. Additional bird sampling should occur at locations where Trichobilharzia szidati were found in snails in order to determine the avian definitive bird host of this parasite in

North Dakota. One previous study in Europe identified teals and mallards as definitive hosts for this species (Rudolfova et al 2005). We expect this to also be the case in North Dakota.

Gigantobilharzia huronensis was the only AS species found in passerine birds. The behavior of Aplexa snails (the intermediate host of G. huronensis) most likely plays an important role in transmission of this parasite to passerines. Aplexa snails were usually found in very shallow areas of water bodies. This promotes prolonged and easy contact between passerine birds and cercariae while birds are searching for food or nesting material. Additional sampling of wetland passerines should reveal more definitive hosts for Gigantobilharzia huronensis.

On at least two occasions, we have found AS with marine life cycles in shorebirds

(marbled godwit and American avocet) in North Dakota. Avian schistosomatids that utilize molluscs from marine environments cannot complete their life cycle in the freshwater bodies of

North Dakota. Therefore, transmission of these AS species could not occur in the state.

The AS diversity seen in North Dakota is due to the high diversity of birds that nest in the region and migrate through it. High avian diversity allows for the introduction and continued maintenance of AS species that may utilize the same intermediate hosts but have the potential to infect multiple bird species. A number of bird species examined during our study were also not found to be infected with avian schistosomatids. These included, for instance, coots, rails,

Northern pintails, mergansers, Ruddy ducks, cormorants and various species of herons. The inability to find AS infected individuals for these species may be due to the low number of individuals examined for each species and the difficulty in finding adult worms in birds when the intensity of infection is low. There were five avian schistosomatid species for which both the intermediate and definitive hosts were identified (Table 7). These bird hosts are all common species in the region with sufficient population densities to allow for completion of the AS life cycle in North Dakota.

Sequence Comparison and Phylogenetic Study of Avian Schistosomatids Found in North Dakota

The nuclear 28s gene is a good target for studies of phylogenetic relationships among the

Schistosomatidae. In addition, the use of this gene made our results compatible and comparable to previous phylogenetic studies of avian schistosomatids (Snyder 2004, Lockyer et al 2003).

However, the relatively low interspecific variability of the 28s gene makes it unsuitable for reliable species differentiation among congeneric species of avian schistosomatids. In order to differentiate between closely related species and produce finer scale intrageneric phylogenies, the high variability in the mitochondrial cox1 gene was also utilized.

Intraspecific variability was practically absent in the 28S gene except for 1-2 nucleotides in one sequence of Trichobilharzia querquedulae and Gigantobilharzia huronensis. In the cox1 gene the intraspecific variability was more pronounced, but was still under 1% while the lowest interspecific variability varied from 5.36% to 14.5%. This allowed for a reliable resolution for differentiation among closely related species, mostly notably we were able to demonstrate the presence of two species of Dendritobilharzia in North America (see below).

Zarowiecki et al (2007) analyzed the mitochondrial genomes of 5 schistosoma species and concluded that both the mitochondrial cox3 and nad5 fragments were better suited for phylogenetic studies of schistosomes as opposed to the commonly used cox1 fragment. Their reasoning was that because these two fragments contain approximately 25% of the variation of the entire mitochondrial genomes, this should then be the most efficient method in capturing genetic variation. In our study, phylogenetic analysis and comparisons required already available sequences from Schistosomatidae taxa. Previous studies on avian schistosomatids have primarily utilized the mitochondrial cox1 barcoding region and so the cox1 partial fragment was sequenced in order to incorporate already available data into the study. Future studies should consider

80 sequencing the recommended mitochondrial cox3 and nad5 fragments that provide better resolution of interspecific/intraspecific levels. Sequencing these genes may help to resolve the phylogenetic relationships among closely related species within the most species-rich

Trichobilharzia clade.

In addition, the nuclear ITS region (ITS1, 5.8S, ITS2) may also provide additional resolution to the relationships between species/genera in the derived avian Schistosomatid clade.

The ITS region exhibits intermediate variability compared to nuclear 28s rDNA (low variability) and mitochondrial cox1 mDNA (high variability).

Phylogenetic Interrelationships among Avian Schistosomatid Genera

Phylogenetic analyses including 28S sequences of all species found in our study combined with sequences of all schistosomatid genera available in the GenBank and using a spirorchid

Hapalotrema mehrai as an outgroup have produced overall well-structured phylogenetic trees with high level of support of most topologies (Figs 22-25). In general, they corresponded well to the previously published phylogenies of the Schistosomatidae (Brant and Loker 2009b, Snyder

2004). We have also conducted separate analyses of alignments that included only the derived avian schistosomatid clade to make them comparable to the analysis presented by Brant et al.

(2013) and Brant and Loker (2013). In both cases (the larger and the smaller alignments) the only difference between the results of Bayesian and Maximum Likelihood analyses was the position of the Allobilharzia-Anserobilharzia clade in relation to the numerous representatives of

Trichobilharzia. The Bayesian analyses consistently resulted in Allobilharzia-Anserobilharzia being nested within one of the sub-clades of the strongly (100%) supported Trichobilharzia clade

(Figs 22 and 26). However, the topology resulted from the Maximum Likelihood analysis was somewhat different. In this case, both the larger and the smaller alignments produced overall

81 weaker supported topologies that showed Allobilharzia-Anserobilharzia as a sister clade to

Trichobilharzia. Brant and Loker (2013, Fig. 1) used essentially the same set of taxa and the same 28S gene region. However, in their Bayesian analysis Allobilharzia-Anserobilharzia appeared as sister group to the clade of Trichobilharzia. We do not have an adequate explanation of this difference. Nevertheless, the unstable position of this clade in different analyses and weak morphological differences between Trichobilharzia, Allobilharzia and Anserobilharzia warrant further detailed consideration of the interrelationships among these digeneans and their taxonomic status.

Trichobilharzia

The pairwise sequence comparison among 7 species of Trichobilharzia found in our study from either snails or birds in North Dakota has demonstrated sequence variability ranging from

0% (between Trichobilharzia sp. 4 and Trichobilharzia sp. A) to 2.74% (between

Trichobilharzia sp. E and Trichobilharzia querquedulae). In some cases only 1-2 nucleotides were different, e.g., between Trichobilharzia querquedulae and Trichobilharzia sp. 4 and

Trichobilharzia sp. A (the latter two species had identical 28S sequences). Overall, the

Trichobilharzia sp. E. was the most divergent species of the genus in our data set (Table 9).

Thus, it can be concluded, that the 28S gene can resolve interspecific differences in most cases, but not all.

The cox1 region utilized in our study proved to be a very useful target for differentiation of closely related congeneric species including all species of Trichobilharzia found in North

Dakota. The levels of interspecific divergence ranged from 5.36% between Trichobilharzia querquedulae and Trichobilharzia sp. 4 to 14.50% between Trichobilharzia sp. E and

Trichobilharzia physellae (Table 9). Interestingly, the levels of sequence divergence observed in

82 the cox1 gene did not fully correspond to the picture provided by the 28S gene. For instance, the pair of species (Trichobilharzia sp. 4 and Trichobilharzia sp. A) that showed no 28S sequence divergence was not the least divergent in the cox1 gene.

Sequences of two Trichobilharzia species found in in our study, namely Trichobilharzia sp.

2 and Trichobilharzia sp. 4, were lacking in the GenBank database and thus represent new genetic lineages available for phylogenetic studies. Their identity requires further clarification.

Considering that the genus Trichobilharzia has been sequenced quite densely in North America it cannot be excluded that at least one of these species-level genetic forms may represent a yet undescribed species.

Dendritobilharzia

Vande Vusse (1980) revised all four formally described Dendritobilharzia species and concluded that all of them should be synonymized with Dendritobilharzia pulverulenta. The results from this study, however, demonstrated the presence of at least two species of

Dendritobilharzia in North America. The variability in the more conserved ribosomal genes did not provide enough resolution for detection/separation of these forms; however, cox1 sequences clearly separated these species (Table 10). One of them (Dendritobilharzia sp. 1) was conspecific with a sequence of Dendritobilharzia cf. pulverulenta obtained from a specimen collected in Europe as well as with the only cox1 sequence of Dendritobilharzia available in the

GenBank database. The other species was represented by several specimens found so far only in

North Dakota. The morphological characteristics that may differentiate between the two species include the location of the cecal bifurcation relative to the anterior testis and the genital pore.

While these characteristics are easy to distinguish they may pose a problem when comparing immature adults. No differences have been identified in female anatomy except for the overall

83 difference in body shape with Dendritobilharzia sp.2 having a more slender body shape compared to the larger and wider Dendritobilharzia sp. 1. It is most likely that Dendritobilharzia sp. 1 represents true Dendritobilharzia pulverulenta while Dendritobilharzia sp. 2 may represent either D. anatinarum described from North America, or a new, yet undescribed, species. Further morphological analysis will clarify this question.

It is also important to identify the snail hosts for Dendritobilharzia in North Dakota.

Previous studies have identified planorbid snails as intermediate hosts for this genus and these snails are also expected to serve as intermediate hosts in North Dakota.

Austrobilharzia

The sequenced specimen of Austrobilharzia showed a significant, species-level divergence in the cox1 gene from sequences of A. variglandis and A. terragalensis available in the

GenBank. Our specimen differed from these species in 13.73% and 17.39% of nucleotide positions, accordingly.

Gigantobilharzia

All nuclear 28s phylogenetic trees produced in this study have shown Gigantobilharzia to be polyphyletic (Figs. 22, 24 and 26). This was also seen in the 28s Bayesian tree of the DAS clade published by Brant and Loker (2013). The polyphyletic nature of this taxon indicates the necessity of a thorough taxonomic revision of Gigantobilharzia and its separation into several independent genera. Further detailed morphological and molecular study of Gigantobilharzia spp. is required to identify the true phylogenetic relationships between current members of this large genus.

Cercarial Morphology

Our morphometric analysis is generally in agreement with the opinion of Horak et al.

(2002) who concluded that cercarial morphological features, including metric characters, in most cases are of limited value for differentiation among species of Trichobilharzia.

To the best of our knowledge, nobody has studied cercariae of T. szidati using SEM.

Among members of Trichobilharzia, only cercariae of T. ocellata were examined using SEM

(Kock and Böckeler 1998; Haas and van de Roemer, 1998). These studies did not focus on the surface morphology other than sensory elements. Only one image in the paper by Kock and

Böckeler (1998) shows spines on the stem of the cercarial tail without any comments in the text.

None of the studies examined furcae or showed the fin fold. Spines on the furcae are documented here for the first time. However, the functional role of the spination of the tail and furcae remains unclear since they cannot assist in attachment to the host or penetration. Spination was also detected on the tail of human schistosomes (Pereira et al. 2013), but they were of very different shape and were arranged in regular rows rather than randomly, as seen in our observation of T. szidati.

Cercarial Dermatitis in North Dakota

Based on this stud , cercarial dermatitis (“swimmer’s itch”) should be considered a concern in the majorit of the state’s water bodies because the most common AS species collected, Trichobilharzia spp. and Gigantobilharzia huronensis, are known to cause dermatitis in people. These two genera had a widespread distribution throughout the state (Fig. 10).

Therefore, any aquatic habitat that provides sufficient resources for snails and birds is a potential source for AS transmission and cercarial dermatitis. The wetlands that were sampled during this study were usually small to moderately sized water bodies. There is a higher risk for AS

85 transmission in small, shallow water bodies, due to the higher concentration of infected snails and lower physical separation between intermediate and definitive hosts. While these small, shallow water bodies may appeal to snails, birds and avian schistosomatid researchers, they are not very likely to be frequented by swimmers or utilized for any other recreational activities that may result in exposure of skin to cercariae. Lakes, reservoirs and other man-made water bodies that are maintained for human recreational activity pose a greater risk of AS transmission.

Monitoring and limiting the number of birds in a location (prohibiting the feeding of ducks) as well as the use of molluscicides are the methods that are being utilized in reducing the risk of swimmer’s itch.

APPENDICES

Appendix A

Map of North Dakota Snail Collection Sites

ed ed

data for each numbered site is available in Appendix B. B. inAppendix is available site numbered each for data record The collected. were snails infected Schistosomatidae where (numbered) sitesall showing Dakota North Mapof 30. Figure

Appendix B

Recorded Data for all Snail Collection Sites

Outwash

Ecoregion

DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains

d, d, numberof snails

CollapsedGlacial Outwash CollapsedGlacial Outwash CollapsedGlacial Outwash CollapsedGlacial

Beach Ridges Beach andSand Deltas

crop crop crop crop crop crop crop crop crop crop crop crop crop crop crop crop crop crop crop crop crop crop crop crop

developed

fallow crop fallow crop fallow

LandCover

deciduousforest deciduousforest deciduousforest

------

7 2

12 94 13 16 64

Aplexa

------

1 6 7 6

Promenetus

------

8 1 1 1 4 5 1

21 21 41 22 15 10

Helisoma

------

1 2 4 3

47 80 12 13

Physa

4 6 3 7 8 5

58 66 47 71 10 48 36 10 12 37 94

285 152 343 235 145 280 151 231 195 290 240 211

Stagnicola

------

2 9 2

63 55 26

166 136 143 106

Lymnaea

13 13 13 13 13 13 13 13 13 13 13

13 13 13 13 13 13 13 13

------

13 13 13 13 13 13 13 13 13 13 13

------

Aug Aug Aug Aug Aug Aug Aug Aug Aug Aug Sep

Date

Jun Jun Jun Jun Jun Jun Jun Jun Jun Jun Jun

Sep Sep Sep Sep Sep Sep Sep Sep

------

7 7 7 7 3 3 5 5 5 5 5

1 1 1 1 1 1 1 1

15 15 15 15 15 31 31 31 31 31 13

ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND

State

99.27

(dec)

100.27 100.84 98.131

97.9003 98.6048 98.4141 98.2966 97.5571 97.8983 98.6061 98.8837 98.8402 99.3346 99.7672 99.8884 99.8994 100.435 100.418 100.941 99.3008 99.1095 99.1066 98.6601 98.7183 101.508 101.518 101.288 99.9694 100.502

- - -

------

Longitude

Recordeddata for snail sites collection from North Dakotaand Minnesota. Data includes GPS coordinates,date was site sample

(dec)

Latitude

48.6164 48.9179 48.9502 46.7721 47.9846

47.44764 47.22569 46.97956 46.70358 46.70219 48.79003 48.82648 48.60907 48.48667 48.90672 48.90695 48.95103 48.92175 48.71944 48.77162 46.95957 47.06646 47.17458 48.16243 48.18011 47.72103 47.98537 47.81918 47.36683 47.47977

1 2 3 4 5 6 7 8 9

10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30

No.

Site

Table20. collected, ecoregion andsurrounding land cover.

Outwash

Drift Plains Drift DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains DriftPlains

Ecoregion

Missou coteau Missou Missoucoteau Missoucoteau Missoucoteau Missoucoteau Missoucoteau Missoucoteau

Glacial Outwash Glacial Outwash Glacial Glacial Missouriplateau

Glacial Lake Glacial Deltas Lake Glacial Deltas

Glacial Lake Lake Glacial Basins

End Moraine End Complex Moraine End Complex Moraine End Complex Moraine End Complex

pasture pasture

LandCover

grasspasture grass grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grass grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture

------

8 3 1

27 16 40 48 40

104

Aplexa

------

5 7 3

Promenetus

------

5 4 7 8 4 8 9 3 2 5

12 12 14 12 10 11

Helisoma

------

3 1 8

28 13 44 13 95

Physa

- - - -

8 5 4 8 9

26 87 93 92 10 71 27 81 10

290 282 128 210 108 240 157 291 128 110 155 200

Stagnicola

------

3 4 4 2 7 8

22 39 56 34 51 22 59 19

155 189 231

Lymnaea

13 13 13 13 13 13 13 13

13 13 13 13 13 13 13 13 13

13 13 13

13 13 13 13 13 13 13 13 13

------

- - -

------

Jul Jul Jul Jul Jul Jul Jul Jul

Jun

Jul Jul Jul Jul Jul Jul Jul Jul Jul

------

Date Jun Jun Jun

Aug Aug Aug Aug Aug Aug Sep Aug Aug

------

- - -

------

2 2 2 2 2 2 2 4 4

6 6 6

6 7 6 6 7 7 7 7 7

16 15 15 15 16 16 17 17

ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND

State

(dec)

97.673 103.97 98.703

102.017 101.771 103.805 101.646 101.632 101.228 97.9252 99.6048 101.812 97.9839 99.7499 98.0205 100.178 102.343 101.965 102.699 97.6648 98.1504 103.257 97.6562 101.556 97.5933 103.248 97.6582 100.522 99.9649 98.5768

- - -

------

Longitude

(dec)

48.2585 47.4507 48.3049 48.6767 48.1671 47.5567

Latitude

48.45842 48.90833 47.37438 46.72672 47.74507 47.62997 48.06485 47.65988 48.61353 47.52067 48.35387 48.75343 47.60037 48.21261 48.77896 48.10423 46.08053 48.76512 47.61543 47.47677 48.97963 48.09047 47.39983 48.13692

39 49 59 31 32 33 34 35 36 37 38 40 41 42 43 44 45 46 47 48 50 51 52 53 54 55 56 57 58 60

able20 cont.

No.

Site

Ecoregion

Missouricoteau Missouricoteau Missouricoteau Missouricoteau Missouricoteau Missouricoteau Missouricoteau Missouricoteau Missouricoteau Missouricoteau Missouricoteau Missouricoteau Missouricoteau Missouricoteau

Missouriplateau Missouriplateau Missouriplateau Missouriplateau

MissouriPlateau

Northern Black Prairie Northern Black Prairie

MissouriCoteau Slope

NorthernMissouri Coteau NorthernMissouri Coteau NorthernMissouri Coteau NorthernMissouri Coteau NorthernMissouri Coteau NorthernMissouri Coteau NorthernMissouri Coteau NorthernMissouri Coteau

LandCover

grasspasture grasspasture grasspasture grasspasture grasspasture grasspasture

herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands

------

26 29

Aplexa

------

1 7

Promenetus

------

3 2 8 2 3 1 8 4 3 3

14 20 17 47 12

Helisoma

------

1 3 1 1 2 3

78 19 80 75

245 227

Physa

------

2 4 3

13 17 11 72 74 53 32 14 44 94 55 47 81 45

254 217 202 105 105 256 116

Stagnicola

------

7 4 8 3

56 12 39 71 46 40 12 25 14 44 72

136 246 149 100 198 110 100 173 109

Lymnaea

13 13 13

13 13

13 13 13 13 13 13 13 13 13 13 13 13 13

13 13 13 13 13 13 13

- - -

- -

13 13 13 13 13

------

- - - - -

Jul Jul Jul Jul Jul Jul Jul Jul Jul Jul Jul Jul Jul

Jun Jun

Sep Sep Aug

Jul Jul Jul Jul Jul

------

Date

- -

Aug Aug Aug Sep Aug Aug Aug

- - -

- - - - -

------

4 7 7 7 7

8 8 8 7 9 9 9

26 17 17 23 23 23 11 23 11 11 26 11 26

22 22

10 13 10

ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND

State

(dec)

103.09 98.552

98.7104 99.9693 98.2279 98.4872 103.206 99.8013 98.5519 98.5519 99.2469 102.634 98.8793 99.1287 98.3577 97.9903 101.175 99.1087 98.7247 99.1173 100.025 98.7894 98.0131 99.1297 98.6754 98.9971 98.6607 99.3171 99.1663

- -

------

98.05465

Longitude

(dec)

46.881

48.1224 48.1754 48.0831 47.2706

Latitude

47.71703 47.26833 48.31005 46.98087 48.08425 46.12825 48.07372 46.63122 45.99395 46.02881 46.86253 48.29638 48.02878 46.46213 46.90987 47.03713 46.02987 48.11505 48.02106 46.02695 48.11597 48.10778 47.75922 48.15147 46.28333

82 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 83 84 85 86 87 88 89 90

No.

Site

Table20 cont.

Ecoregion

Saline Saline Area

RiverBreaks

TurtleMountains TurtleMountains TurtleMountains TurtleMountains TurtleMountains TurtleMountains

Northern Black Prairie Northern Black Prairie Northern Black Prairie Northern Black Prairie

Tewaukon/Big Stagnation Stone Moraine Tewaukon/Big Stagnation Stone Moraine Tewaukon/Big Stagnation Stone Moraine

open open water openwater openwater openwater openwater openwater openwater openwater openwater openwater

LandCover

herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands herbaceouswetlands

------

170

Aplexa

------

Promenetus

------

8 8 6 8 1

12 12 17

Helisoma

------

4 5 2 3 4

11 29

Physa

- - - -

95 91 53 38 10

101 458 165 458 112 396 174 138 555 171

Stagnicola

------

1 1 2 2

97 81 23 88 14

156 100 105 120 103

Lymnaea

13 13 13

13 13 13 13 13 13 13 13 13 13

- 13 13 13 13 13

- - -

------

- - - - -

Jul Jul Jul Jul Jul Jul Jul Jul Jul Jul

Jun

Aug Aug Aug

May

------

Date

Sep Aug Sep Sep Sep

- - -

- - - - -

6 3 7 7 7

11 21 11 26 26 16 21 21 27 27

10 10 10

ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND

MN MN MN MN MN

State

(dec)

98.0117 95.2242 98.7029 98.1831 98.1831 99.3128 99.1291 97.2399 97.7374 99.1215 98.3705 98.2019 96.9499 98.3682 97.9723 100.743 94.9667 94.9717 94.9376 94.4986

------

Longitude

(dec)

47.19

47.255

48.2095

Latitude

48.07893 48.12605 47.90406 47.90406 47.19318 46.47145 47.81408 46.10317 47.10695 46.00847 46.45537 45.97768 46.10975 47.48748 47.05867 47.40017 47.10604

96 91 92 93 94 95 97 98 99

No.

106 100 101 102 103 104 105 107 108 109 110

Site

Table20 cont.

Appendix C Data on Snails Infected with Schistosomatidae Cercariae

Table 21. Data for schistosomatidae infected snails including snail host genus, AS species and sequenced genes. Site Snail Snail State Date Ext No. AS species 28s Cox1 No. No. genus Collected 4 2410 Stagnicola ND 22-Aug-13 2754neo Trichobilharzia Y N 4 2403 Lymnaea ND 15-Aug-13 2747neo Trichobilharzia szidati Y N 4 2400 Lymnaea ND 15-Aug-13 2744neo Trichobilharzia szidati Y Y 4 2401 Lymnaea ND 15-Aug-13 2745neo Trichobilharzia szidati Y Y 4 2405 Lymnaea ND 17-Aug-13 2749neo Trichobilharzia szidati Y Y 4 2406 Lymnaea ND 18-Aug-13 2750neo Trichobilharzia szidati Y Y 4 2407 Lymnaea ND 19-Aug-13 2751neo Trichobilharzia szidati Y N 4 2408 Lymnaea ND 20-Aug-13 2752neo Trichobilharzia szidati Y N 4 2409 Lymnaea ND 21-Aug-13 2753neo Trichobilharzia szidati Y Y 7 2423 Lymnaea ND 31-Aug-13 2755neo Trichobilharzia szidati Y N 8 2425 Stagnicola ND 31-Aug-13 2505neo Trichobilharzia sp E Y Y 9 2428 Stagnicola ND 31-Aug-13 2780neo Trichobilharzia sp 4 Y Y 9 2427 Stagnicola ND 31-Aug-13 2509neo Trichobilharzia szidati Y Y 9 2426 Stagnicola ND 31-Aug-13 2889neo Trichobilharzia sp E Y Y 10 2429 Stagnicola ND 31-Aug-13 2507neo Trichobilharzia Y N 11 2430 Stagnicola ND 1-Sep-13 2495neo Schistosomatium douthitti Y N 11 2431 Stagnicola ND 1-Sep-13 2781neo Schistosomatium douthitti Y N 14 2433 Lymnaea ND 1-Sep-13 2506neo Schistosomatium douthitti Y N 14 2432 Lymnaea ND 1-Sep-13 2498neo Trichobilharzia szidati Y Y 16 2435 Lymnaea ND 1-Sep-13 2782neo Schistosomatium douthitti Y N 16 2434 Lymnaea ND 1-Sep-13 2502neo Trichobilharzia szidati Y Y 18 2498 Stagnicola ND 1-Sep-13 2508neo Trichobilharzia sp E Y Y 24 2553 Lymnaea ND 13-Sep-13 2888neo Trichobilharzia sp A Y Y 25 1215 Lymnaea ND 3-Jun-13 2777neo Trichobilharzia szidati Y N 34 1229 Aplexa ND 2-Jul-13 2469neo Gigantobilharzia huronensis Y Y 34 1230 Aplexa ND 2-Jul-13 2473neo Gigantobilharzia huronensis Y Y 34 1234 Aplexa ND 2-Jul-13 2474neo Gigantobilharzia huronensis Y Y 34 1231 Physa ND 2-Jul-13 2472neo Gigantobilharzia huronensis Y Y 35 1597 Lymnaea ND 15-Jul-13 2894neo Trichobilharzia szidati Y Y 35 1598 Lymnaea ND 15-Jul-13 2567neo Trichobilharzia szidati Y Y 37 1238 Stagnicola ND 2-Jul-13 2512neo Trichobilharzia sp 4 Y Y 38 1599 Lymnaea ND 15-Jul-13 2568neo Schistosomatium douthitti Y N 38 1601 Lymnaea ND 15-Jul-13 2570neo Trichobilharzia sp E Y Y 38 1600 Lymnaea ND 15-Jul-13 2569neo Trichobilharzia szidati Y Y 39 2129 Aplexa ND 6-Aug-13 2613neo Gigantobilharzia huronensis Y Y 39 2131 Aplexa ND 6-Aug-13 2778neo Gigantobilharzia huronensis Y Y 39 2143 Lymnaea ND 6-Aug-13 2619neo Schistosomatium douthitti Y N 39 2142 Stagnicola ND 6-Aug-13 2618neo Schistosomatium douthitti Y N 39 2126 Aplexa ND 6-Aug-13 2611neo Trichobilharzia querquedulae Y Y 39 2128 Aplexa ND 6-Aug-13 2612neo Trichobilharzia querquedulae Y Y 39 2130 Aplexa ND 6-Aug-13 2614neo Trichobilharzia querquedulae Y N 39 2137 Stagnicola ND 6-Aug-13 2617neo Trichobilharzia sp E Y Y

Table 21 cont. Site Snail Snail State Date Ext No. AS species 28s Cox1 No. No. genus Collected 41 1602 Aplexa ND 15-Jul-13 2571neo Gigantobilharzia huronensis Y Y 42 2123 Lymnaea ND 6-Aug-13 2609neo Trichobilharzia szidati Y Y 42 2124 Lymnaea ND 6-Aug-13 2610neo Trichobilharzia szidati Y N 44 1826 Stagnicola ND 16-Jul-13 2595neo Trichobilharzia sp A Y Y 44 1619 Stagnicola ND 16-Jul-13 2573neo Trichobilharzia sp E Y Y 44 1618 Stagnicola ND 16-Jul-13 2945neo Trichobilharzia sp E Y Y 45 2159 Lymnaea ND 7-Aug-13 2620neo Trichobilharzia szidati Y Y 45 2160 Lymnaea ND 7-Aug-13 2621neo Trichobilharzia szidati Y N 45 2163 Lymnaea ND 7-Aug-13 2623neo Trichobilharzia szidati Y Y 45 2165 Lymnaea ND 7-Aug-13 2953neo Trichobilharzia szidati Y N 49 1624 Aplexa ND 16-Jul-13 2576neo Gigantobilharzia huronensis Y Y 49 1625 Aplexa ND 16-Jul-13 2582neo Gigantobilharzia huronensis Y Y 49 1627 Aplexa ND 16-Jul-13 2584neo Gigantobilharzia huronensis Y Y 49 1634 Aplexa ND 16-Jul-13 2591neo Gigantobilharzia huronensis Y Y 49 1635 Aplexa ND 16-Jul-13 2592neo Gigantobilharzia huronensis Y Y 49 1636 Aplexa ND 16-Jul-13 2593neo Gigantobilharzia huronensis Y Y 49 1626 Aplexa ND 16-Jul-13 2947neo Gigantobilharzia huronensis Y Y 49 1623 Stagnicola ND 16-Jul-13 2575neo Trichobilharzia sp 4 Y Y 49 1628 Aplexa ND 16-Jul-13 2585neo Trichobilharzia querquedulae Y N 49 1622 Stagnicola ND 16-Jul-13 2574neo Trichobilharzia sp A Y Y 49 1629 Stagnicola ND 16-Jul-13 2586neo Trichobilharzia sp A Y Y 50 1274 Aplexa ND 2-Jul-13 2616neo Gigantobilharzia huronensis Y Y 53 2176 Lymnaea ND 7-Aug-13 2627neo Trichobilharzia szidati N N 53 2177 Lymnaea ND 7-Aug-13 2628neo Trichobilharzia szidati Y Y 53 2178 Lymnaea ND 7-Aug-13 2661neo Trichobilharzia szidati Y Y 53 2180 Lymnaea ND 7-Aug-13 2662neo Trichobilharzia szidati Y N 53 2181 Lymnaea ND 7-Aug-13 2663neo Trichobilharzia szidati Y N 53 2182 Lymnaea ND 7-Aug-13 2664neo Trichobilharzia szidati Y Y 53 2183 Lymnaea ND 7-Aug-13 2665neo Trichobilharzia szidati Y N 53 2184 Lymnaea ND 7-Aug-13 2666neo Trichobilharzia szidati Y Y 53 2185 Lymnaea ND 7-Aug-13 2667neo Trichobilharzia szidati Y Y 53 2186 Lymnaea ND 7-Aug-13 2668neo Trichobilharzia szidati Y Y 53 2188 Lymnaea ND 7-Aug-13 2687neo Trichobilharzia szidati Y Y 53 2174 Stagnicola ND 7-Aug-13 2626neo Trichobilharzia szidati Y Y 57 1292 Stagnicola ND 4-Jul-13 2615neo Trichobilharzia sp 4 Y Y 61 1630 Aplexa ND 17-Jul-13 2587neo Gigantobilharzia huronensis Y N 61 1631 Aplexa ND 17-Jul-13 2588neo Gigantobilharzia huronensis Y N 62 1216 Stagnicola ND 22-Jun-13 2470neo Trichobilharzia sp 4 Y Y 65 1632 Lymnaea ND 17-Jul-13 2589neo Trichobilharzia szidati Y Y 65 1633 Lymnaea ND 17-Jul-13 2590neo Trichobilharzia szidati Y Y 68 1405 Lymnaea ND 7-Jul-13 2531neo Trichobilharzia szidati Y N 68 1407 Lymnaea ND 7-Jul-13 2533neo Trichobilharzia szidati Y Y 68 1408 Lymnaea ND 7-Jul-13 2534neo Trichobilharzia szidati Y Y 68 1436 Lymnaea ND 7-Jul-13 2898neo Trichobilharzia szidati Y Y 68 1440 Lymnaea ND 7-Jul-13 2900neo Trichobilharzia szidati Y Y 68 1396 Lymnaea ND 7-Jul-13 2901neo Trichobilharzia szidati Y Y 68 1406 Lymnaea ND 7-Jul-13 2902neo Trichobilharzia szidati Y Y

Table 21 cont. Site Snail Snail State Date Ext No. AS species 28s Cox1 No. No. genus Collected 68 1438 Lymnaea ND 7-Jul-13 2899neo Trichobilharzia szidati Y Y 69 1760 Lymnaea ND 23-Jul-13 2594eno Schistosomatium douthitti Y N 72 2504 Lymnaea ND 7-Sep-13 2906neo Trichobilharzia szidati Y Y 72 2505 Lymnaea ND 7-Sep-13 2504neo Trichobilharzia szidati Y N 75 2509 Lymnaea ND 10-Sep-13 2887neo Trichobilharzia szidati Y N 77 1833 Lymnaea ND 23-Jul-13 2597neo Trichobilharzia szidati Y Y 77 1834 Lymnaea ND 23-Jul-13 2954neo Trichobilharzia szidati Y N 78 1443 Lymnaea ND 11-Jul-13 2552neo Trichobilharzia szidati Y N 78 1444 Lymnaea ND 11-Jul-13 2553neo Trichobilharzia szidati Y Y 78 1445 Lymnaea ND 11-Jul-13 2554neo Trichobilharzia szidati Y Y 78 1447 Lymnaea ND 11-Jul-13 2555neo Trichobilharzia szidati Y Y 78 1448 Lymnaea ND 11-Jul-13 2560neo Trichobilharzia szidati Y Y 81 1449 Lymnaea ND 11-Jul-13 2561neo Trichobilharzia szidati Y N 82 2017 Physa ND 26-Jul-13 2948neo Gigantobilharzia huronensis Y N 82 2016 Lymnaea ND 26-Jul-13 2601neo Schistosomatium douthitti Y N 82 1957 Lymnaea ND 26-Jul-13 2600neo Trichobilharzia szidati Y Y 83 2517 Stagnicola ND 13-Sep-13 2494neo Trichobilharzia Y N 90 1956 Lymnaea ND 26-Jul-13 2599neo Trichobilharzia szidati Y Y 91 1452 Aplexa ND 11-Jul-13 2893neo Gigantobilharzia huronensis Y Y 91 1451 Stagnicola ND 11-Jul-13 2563neo Trichobilharzia Y N 91 1450 Stagnicola ND 11-Jul-13 2562neo Trichobilharzia sp A Y Y 92 2508 Lymnaea ND 6-Sep-13 2783neo Schistosomatium douthitti Y N 93 1183 Lymnaea ND 29-May-13 2471neo Trichobilharzia szidati Y Y 93 1184 Lymnaea ND 29-May-13 2776neo Trichobilharzia szidati Y Y 94 2020 Lymnaea ND 26-Jul-13 2907neo Schistosomatium douthitti Y N 95 2281 Stagnicola ND 10-Aug-13 2693neo Trichobilharzia Y N 95 2284 Stagnicola ND 10-Aug-13 2723neo Trichobilharzia Y N 95 2279 Stagnicola ND 10-Aug-13 2692neo Trichobilharzia Y N 95 2287 Stagnicola ND 10-Aug-13 2725neo Trichobilharzia sp 2 Y Y 95 2283 Stagnicola ND 10-Aug-13 2946neo Trichobilharzia sp 2 Y Y 95 2282 Stagnicola ND 10-Aug-13 2694neo Trichobilharzia sp A Y Y 95 2286 Stagnicola ND 10-Aug-13 2724neo Trichobilharzia sp A Y Y 95 2288 Stagnicola ND 10-Aug-13 2726neo Trichobilharzia sp A Y Y 95 2292 Stagnicola ND 10-Aug-13 2728neo Trichobilharzia sp A Y Y 95 2293 Stagnicola ND 10-Aug-13 2729neo Trichobilharzia sp A Y Y 95 2294 Stagnicola ND 10-Aug-13 2730neo Trichobilharzia sp A Y Y 95 2295 Stagnicola ND 10-Aug-13 2731neo Trichobilharzia sp A Y Y 95 2275 Stagnicola ND 10-Aug-13 2779neo Trichobilharzia sp A Y Y 95 2290 Stagnicola ND 10-Aug-13 2727neo Trichobilharzia sp E Y N 95 2272 Lymnaea ND 10-Aug-13 2688neo Trichobilharzia szidati Y N 95 2273 Lymnaea ND 10-Aug-13 2689neo Trichobilharzia szidati Y Y 95 2274 Lymnaea ND 10-Aug-13 2690neo Trichobilharzia szidati Y Y 97 2092 Lymnaea ND 3-Aug-13 2608neo Trichobilharzia szidati Y Y 98 2506 Stagnicola ND 7-Sep-13 2500neo Trichobilharzia Y N 98 2507 Stagnicola ND 7-Sep-13 2905neo Trichobilharzia sp 4 Y Y 99 2018 Lymnaea ND 26-Jul-13 2603neo Trichobilharzia szidati Y N 99 2019 Lymnaea ND 26-Jul-13 2604neo Trichobilharzia szidati Y Y

Table 21 cont. Site Snail Snail State Date Ext No. AS species 28s Cox1 No. No. genus Collected 101 2303 Lymnaea ND 10-Aug-13 2738neo Schistosomatium douthitti Y N 101 2306 Stagnicola ND 10-Aug-13 2741neo Trichobilharzia sp A Y Y 101 2297 Lymnaea ND 10-Aug-13 2732neo Trichobilharzia szidati Y N 101 2298 Lymnaea ND 10-Aug-13 2733neo Trichobilharzia szidati Y N 101 2304 Lymnaea ND 10-Aug-13 2739neo Trichobilharzia szidati Y N 101 2305 Lymnaea ND 10-Aug-13 2740neo Trichobilharzia szidati Y Y 101 2302 Lymnaea ND 10-Aug-13 2890neo Trichobilharzia szidati Y N 101 2301 Lymnaea ND 10-Aug-13 2891neo Trichobilharzia szidati Y Y 101 2299 Lymnaea ND 10-Aug-13 2903neo Trichobilharzia szidati Y Y 101 2300 Lymnaea ND 10-Aug-13 2904neo Trichobilharzia szidati Y Y 102 2307 Stagnicola ND 10-Aug-13 2742neo Schistosomatium douthitti Y N 108 1827 Physa MN 21-Jul-13 2596neo Gigantobilharzia huronensis Y Y 109 2021 Lymnaea MN 27-Jul-13 2606neo Trichobilharzia szidati Y Y 109 2022 Lymnaea MN 27-Jul-13 2607neo Trichobilharzia szidati Y Y

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